Cxcl12-null embryos have dysplastic, misaligned, and hyperplastic semilunar valves (SLVs). In this study, we show that CXCL12 signaling via its receptor CXCR4 fulfills distinct roles at different stages of SLV development, acting initially as a guidance cue to pattern cellular distribution within the valve primordia during the endocardial-to-mesenchymal transition (endoMT) phase and later regulating mesenchymal cell proliferation during SLV remodeling. Transient, anteriorly localized puncta of internalized CXCR4 are observed in cells undergoing endoMT. In vitro, CXCR4+ cell orientation in response to CXCL12 requires phosphatidylinositol 3-kinase (PI3K) signaling and is inhibited by suppression of endocytosis. This dynamic intracellular localization of CXCR4 during SLV development is related to CXCL12 availability, potentially enabling activation of divergent downstream signaling pathways at key developmental stages. Importantly, Cxcr7-/- mutants display evidence of excessive CXCL12 signaling, indicating a likely role for atypical chemokine receptor CXCR7 in regulating ligand bioavailability and thus CXCR4 signaling output during SLV morphogenesis.  •Cxcl12-null miceSemilunar valves are dysplastic, misaligned, and hyperplastic in •Cells undergoing endoMT express punctate CXCR4 and require CXCL12 for guidance•CXCL12 regulates proliferation of valve mesenchymal cells after E12.5•CXCR7 is required as a sink for CXCL12 during valve development in vivo and in vitro analysis, Ridge et al. show that the CXCL12/CXCR4/CXCR7 signaling axis fulfills distinct roles at different stages of mouse semilunar valve development. Regulation of ligand availability and CXCR4 activity by CXCR7 may be key to enabling alternative functional outcomes from chemokine signaling within a single tissue.Using  The semilunar valves (SLVs) are positioned at the base of the great arteries and ensure unidirectional blood flow . The SLVCxcl12-ablated mouse embryos /AKT and mitogen-activated protein kinase (MAPK); this is followed by rapid phosphorylation of CXCR4 at its C terminus (by GPCR-specific kinases) and β-arrestin-mediated internalization , represent the most prevalent class of congenital heart malformations, affecting 1%–5% newborns . Moreove embryos . The che embryos . As well embryos , CXCL12  embryos . Bindinglization . As partlization . In addilization . A seconlization . CXCR7 clization .+ cells undergoing endoMT, a phase associated with the presence of punctate, internalized CXCR4. Furthermore, the ability of CXCR4+ OFT MCs to respond to CXCL12 in vitro depends on both PI3K signaling and endocytosis. Hyperplasia observed in NC-specific knockouts of CXCR4 indicates the involvement of further cell lineages in SLV remodeling post-endoMT. Finally, analysis of Cxcr7 null mutants and ex vivo receptor inhibition experiments suggest that CXCR7 may act as a chemokine scavenger to modulate CXCL12 levels at different stages of SLV development.Although CXCL12, CXCR4, and CXCR7 are all known to be involved in SLV formation, mechanistic details remain unclear. Here, we demonstrate that CXCL12 signaling affects distinct aspects of SLV morphogenesis, with an early requirement for CXCL12 signaling in regulating leaflet shape and alignment and a later role in downregulating the proliferation of valve mesenchymal cells (MCs). Analysis of mutant mouse embryos showed that CXCL12 provides a directional cue for CXCR4Cxcl12-null mouse embryos  at E18.5  to wild-type/Cxcl12+/− controls (n = 18) at E15.5 revealed dysplasia  in 100% of null SLVs and misalignment (of left and right leaflets) in 59.4% . Dysplasia and subtle misalignment of the valve leaflets was also observed earlier, at E12.5 . 3D reconstruction of E15.5 SLVs enabled measurement of leaflet volumes, revealing variable hyperplasia in Cxcl12 nulls , although this was only significant for the PV  were more evenly distributed along the OFT  compared to proximal OFT (40.1%) and E12.5 leaflets (10.6%)  were detected in ECs (E10.5 and E11.5) and MCs  H–2Hii. NCxcl12 expression in the OFT was broadly complementary to CXCR4, as determined by GFP fluorescence in Cxcl12GFP/+ hearts. In addition to high expression throughout the OFT wall, CXCL12-GFP was detected at lower levels in the cushions at E10.5–E11.5, declining in a distal-proximal fashion  cells found in Tie2-Cre;R26RYFP/+ OFT were GFP+, showing that they derived from endocardium . We observed displacement of CXCR4+ MCs from lateral to central OFT cushion, as well as a significant shift in localization from distal to proximal in Cxcl12 nulls  compared to wild-type controls , suggesting that CXCL12 secreted by the OFT is required for cell guidance during endoMT. Interestingly, co-localization of CXCR4 puncta with the Golgi and a further anterior marker, the centrosome, was observed in both MCs and ECs  polarity in response to external cues. To test whether direction sensing was adversely affected in CXCR4r marker . Numbersr marker R–3T. The and ECs N–S3N”’.+ cells orientated away from the gel surface (and thus toward the gradient) from 21.9%  to 28.3% of total cells, i.e., a 1.29-fold increase  or PI3K/AKT  signaling cascades prior to application of the CXCL12 gradient. Whereas inhibition of MEK1/2 had no significant effect on the orientation of CXCL12-treated cells, inhibition of PI3K completely abolished the response to CXCL12 C. Treatm+ cells is the earliest manifestation of the Cxcl12 null phenotype; however, null SLVs also exhibit hyperplasia  and hyperplasia (E13.5 onward) indicates distinct roles of CXCL12 signaling at different stages of SLV development.Mispositioning of CXCR4erplasia M and 1N.a , proliferation of CXCR4+ MCs was not significantly reduced in Cxcl12 nulls at E15.5. In contrast, proliferation of CXCR4− MCs was significantly decreased in both controls (56% in AoV and 69% in PV) and Cxcl12 nulls (43.2% in the AoV and 51% in the PV). Proliferation rates in endocardial cells were similar in controls and nulls , whereas 13/13 Pax3-Cre conditional mutants exhibited a milder, hyperplastic phenotype characterized by thickening of the hinge regions, without misalignment of the leaflets  and Pax3-Cre  conditional nulls , leaflet alignment was normal in all Pax3-Cre;Cxcr4fl/+ SLVs.To compare the roles of CXCR4 in SLV cells of differing origins, CXCR4 was conditionally ablated in endocardial and NC lineages using al nulls I. HoweveTie2-Cre conditional null hearts only . These results are consistent with an early role for CXCR4 in the positioning of endocardially derived MCs. Analysis of CXCR4 expression at E13.5 showed a comparable decrease in both types of conditional null , showing that hyperplasia manifests between E13.5 and E15.5. Thus, whereas CXCR4 expressed in endocardial derivatives is required for the process of leaflet shaping and alignment between E11.5 and E12.5, CXCR4+ cells of both endocardial and NC origin contribute to the regulation of leaflet growth at later developmental stages.At E12.5, misaligned, malformed SLVs were observed in rts only T–5Y. Abnnal null J–S4Q’. PCxcr7 mutants  and Cxcl12 (n = 6) nulls, compared with controls  showed evidence of leaflet misalignment  appeared somewhat dysplastic  were milder.CXCR7 can act synergistically, antagonistically, or independently of CXCR4  . Both pu compare L–S5L”. OCxcr7 nulls revealed that CXCR4 puncta persist after E11.5, with a concomitant reduction in CXCR4 located on the cell surface . At E11.5, speckled b-CXCL12 signal was detected predominantly in ECs  or CXCR4  and SLVs (E12.5 and e15.5) in order to assess the level of receptor activation in the absence of CXCR7 , E12.5  and E15.5 , i.e., more than double the increases seen in wild types  in wild types, the increased proportion of phos-CXCR4 at E11.5 compared to E12.5 and E15.5, combined with the relative frequency of CXCR4 puncta at E11.5 and E12.5 (see + cells are exposed to a higher level of CXCL12 at E11.5 than at later stages of SLV development. (2) Increased internalization and activation of CXCR4 in the absence of CXCR7 indicate exposure to excess CXCL12, suggesting that sequestration by CXCR7 normally occurs from E12.5 onward. Thus, scavenging of CXCL12 by CXCR7 within SLV primordia may be important for modulating CXCL12-activated pathways during valve development.Phosphorylation of CXCR4 by CXCL12 at multiple C-terminal sites occurs rapidly after activation , and expof CXCR7 E–S6N’. Asections E–S6N’. Isections H. This ild types H. Thus, 12.5 see I implies+ MCs. Sequestration of CXCL12 by CXCR7 modulates the level of cellular exposure to ligand during these temporally separated processes  signaling , epidermal growth factor (EGF), and NF1/RAS signaling pathways have all been implicated in driving expansion of endocardial cushion mesenchyme from E11.5  in wild types coincides with the expansion of SLV mesenchymal CXCR7 expression between E11.5 and E12.5; thus, we surmise that CXCR7 normally sequesters CXCL12 in the SLVs from E12.5 onward. CXCL12 sequestration by CXCR7 is reminiscent of previous studies on several tissues, including mammalian interneurons and zebrafish primordial germ cells and lateral line primordium .Requests for further information should be directed to the lead contact, Sarah Ivins (Cxcr7flox/flox and Cxcr7+/− lines (latter derived from the former) due to the presence of an MTA from https://www.mmrrc.org/, and the Cxcl12GFP/+ line to an MTA from Frontier Medical Sciences Kyoto University (contact Prof Takashi Nagasawa).This study did not generate new unique reagents. There are restrictions to the availability of the All animal work was undertaken in accordance with UK Home Office regulations. Individual mouse lines are listed in the 2 in Dulbecco’s Modified Eagle Medium (DMEM)  supplemented with 10% fetal bovine serum (FBS)  and penicillin/streptomycin  unless otherwise stated. OFTs from both male and female embryos were utilized and grouped for downstream treatment based on genotype alone.Pregnant females were sacrificed by cervical dislocation and embryos were harvested at E10.5. Dissected outflow tract explants were grown at 37°C, 5% COAll protocols were performed at room temperature, unless otherwise stated.2O . PCRs were set up with BIOTAQ , Amplitaq Gold 360  or KAPA2G Taq  polymerase kits, as per the manufacturer’s instructions. Genotyping primer pairs are detailed in DNA samples for genotyping were extracted and purified from embryonic yolk sacs (E ≤ 11.5) or tail clippings (E ≥ 12.5) using standard methods. Briefly, tissue was digested overnight at 56°C in 400 μL lysis buffer , 10mM TRIS , 10mM EDTA , 0.5% SDS , 400ug/mL proteinase K ), and DNA was precipitated by addition of an equal volume of isopropanol . After centrifugation , DNA pellets were washed with 70% (v/v) ethanol , and then resuspended in an appropriate volume of ddHWhole-mount hearts were fixed in 4% paraformaldehyde (PFA)  for 15-30 minutes. For tissue sections, hearts were fixed in 4% PFA for 15-30 minutes (for cryosections) or overnight (for paraffin sections). All specimens were subsequently thoroughly washed with PBS to remove excess PFA. To prepare samples for paraffin embedding, samples were dehydrated through an increasing ethanol concentration series (50/70/85/95/100% (v/v) ethanol in PBS), washed twice with 100% Histo-Clear , and then moved to 50/50 Histo-Clear/paraffin at 65°C. Samples were then washed with at least three changes of 100% paraffin . All incubations were for 30-60 minutes. For cryo-embedding, samples were first incubated overnight at 4°C in 30% (w/v) sucrose , The following day, samples were incubated in 50/50 30% sucrose/optical cutting temperature compound (OCT)  for 30 minutes, followed by 3 washes in 100% OCT; samples were then snap-frozen in 100% OCT.2O, 2 minutes each) and washed in dH2O for 2 × 5 minutes. For H&E staining, slides were incubated in hematoxylin  for 3 minutes and then rinsed in at least 3 changes of tap water for a total of ten minutes. Next, slides were incubated in eosin  for 90 s, washed in ddH2O for 2 minutes, and allowed to dry overnight. The next day, coverslips were mounted using DPX mounting media .Paraffin sections (10μ) were dewaxed in Histo-Clear (2 × 10 minutes), then rehydrated through a decreasing ethanol gradient . Briefly, sections were incubated in alcian blue solution (pH2.5) for 30 minutes and counter stained with nuclear fast red, as per the manufacturer’s instructions. Sections were imaged using a Zeiss Axioplan2 Brightfield microscope with Zeiss Axiocam HRc color camera.E15.5 hearts were permeabilised in PBS with 0.1% (v/v) Tween 20  (PBST), blocked for one hour in PBST with 10% goat serum , and incubated in anti-PECAM-1 primary antibody see  diluted https://imagej.net/software/fiji ) and processed for analysis and publication. N.B. CXCR4 antibody clones UMB2 and EPUMBR3 were used interchangeably to detect unphosphorylated CXCR4; clones 247506 and 2B11 detect total CXCR4. Unless otherwise stated in the Figure Legends, anti-CXCR4 antibody refers to clones UMB2 or EPUMBR3.Immunolabelling was carried out using standard protocols. Briefly, frozen tissue sections (10μ) were permeabilised in 0.5% (v/v) Triton X-100  for 5 minutes before blocking , 0.1% (v/v) Triton X-100) for 1 hour. Tissue sections were incubated in primary antibody diluted in blocking buffer overnight at 4°C. Antibodies see  were useSLV leaflet alignment was determined by examining consecutive coronal sections or z stack images . As alignment of the non-coronal or anterior leaflets in relation to the other leaflets was more likely to be obscured by skewing of the section or imaging plane, alignment of the left and right leaflets only was assessed in order to avoid misleading results. Leaflets were counted as misaligned when the bottom and top edges of the base of one leaflet were displaced by more than half the width of the base of the opposing leaflet in at least 3 consecutive sections.Frozen tissue sections were immunostained with anti-GM130 , anti-CXCR4 (clone UMB2) and anti-PECAM-1 antibodies, as described in ‘Cxcl12 nulls and controls, 3-6 sections from 4 embryos of each genotype were analyzed;For Cxcr7 nulls and controls, 4-6 sections from 3 embryos per genotype were analyze.for OFT explant cultures were prepared using standard protocols with some modifications . BrieflyFor immunostaining, slides were treated as described in the ‘+ mesenchymal (PECAM-1-) cells, ii) total CXCR4- mesenchymal cells (PECAM-1-) or iii) total endocardial (PECAM-1+) cells within the semilunar valve leaflets. 4 Cxcl12 nulls were compared to 4 controls (wild-type/heterozygous) at E12.5, and 4 Cxcl12 nulls were compared to 5 controls (wild-type/heterozygous) at E15.5; 4-6 sections were analyzed per embryo.EdU (5-ethynyl-2'-deoxyuridine) incorporation experiments were conducted using the Click-iT EdU Cell Proliferation kit  and detected as per the manufacturer’s instructions. Briefly, pregnant females were injected with 200 μL 10uM EdU (in PBS) at the relevant developmental stage. Embryos were harvested 75 mins later and processed for immunostaining with anti-PECAM-1 and anti-unphosphorylated CXCR4 antibodies, as previously described in ‘Hearts from wild-type E11.5 or E12.5 embryos were dissected into PBS . Following a brief incubation in PBS + 2% FBS at 37°C, tissue was blocked in DMEM containing 3.78uM unlabelled streptavidin  for 30 minutes at 37°C. Samples were washed twice in PBS + 2% FBS, and then incubated in media containing 200nM biotinylated-CXCL12  conjugated to streptavidin-594  for 30 minutes at 37°C. The biotinylated-CXCL12: streptavidin-594 conjugate was formed by incubating these components in a 1:4 molar ratio at room temperature for 3 minutes immediately prior to addition to the tissue, as per manufacturer’s instructions. For competition experiments, samples were co-incubated with 2μM unlabelled CXCL12 . Samples were washed twice in ice cold DMEM, blocked with PBS + 0.5% BSA for 5 minutes at 4°C, fixed with 4% PFA for 10 minutes, and then washed twice with PBS at 4°C. Specimens were then embedded in OCT for cryosectioning and immunohistochemistry with anti-PECAM-1 antibody as described in ‘General Immunohistochemistry and Quantification’ with the exception that blocking was performed using PBS + 2%BSA. Images were acquired on a spinning disk microscope, as detailed above. Images were exported to FIJI and processed for publication.Hearts were dissected from E12.5 wild-type embryos into pre-warmed PBS, then briefly incubated in 1ml PBS + 2% FBS at 37°C. The hearts were then placed into Eppendorf tubes containing 200 μl DMEM supplemented with inhibitor  according to the manufacturer’s instructions. Sections were then washed, blocked and immunostained using anti-PECAM-1 and anti-unphosphorylated CXCR4 antibodies (clone EPUMBR3), and imaged as described in ‘http://theolb.readthedocs.io/en/latest):Serial frozen tissue sections were incubated at room temperature for 3 hours in the presence or absence of lambda protein phosphatase (800u/slide)  diluted in the supplied buffer  from stage-matched E11.5 wild-type and mutant embryos (Cxcl12−/− and Cxcr7−/−) and presented as a percentage of total CXCR4+ MCs. Sections were stained with anti-PECAM-1 and anti-CXCR4 (clone EPUMBR3) antibodies and DAPI, and CXCR4+/PECAM-1- cells were counted using the Fiji Cell Counter plugin. OFT sectors were determined by dividing the OFT  into roughly equal thirds, and using OFT morphology to fine-tune the sector boundaries. Equal numbers of sections (4-5) were analyzed per sector, ensuring also that the sections used for analysis were spread evenly along the length of the individual sector. 4 stage-matched controls were compared to 5 nulls for both Cxcl12−/− and Cxcr7−/− OFTs.Numbers of CXCR4+ MCs and ECs and their distribution within the OFT  per sector. Equal numbers of sections (4-5) were analyzed per sector, as above. For this comparison, distal and medial sector cell counts were merged to give a ‘valve-forming region’ count. For E12.5 hearts (n = 4), 5-6 sections per SLV were analyzed.Sections from wild-type E11.5 or E12.5 hearts were stained with anti-PECAM-1 and anti-CXCR4 (clone EPUMBR3) antibodies and DAPI. For E11.5 hearts (n = 4), OFT sectors  were determined as above. Numbers of CXCR4Cxcl12−/− or Cxcr7−/−) and pooled littermate controls (wild-type or heterozygous mutant) were matched for dorso-ventral position within the SLV. 9-12 coronal sections per SLV were outlined and measured in Fiji; total leaflet area was measured for each section and mean values calculated for controls and nulls. A total of 10 controls, 8 Cxcr7−/− and 6 Cxcl12−/− were analyzed.Cryosections of E15.5 hearts were stained with anti-PECAM-1 antibody and DAPI. Where possible, for comparison purposes, sections from null (All other procedures used for quantification are provided in the relevant section of https://www.graphpad.com). All data presented in histograms are presented as mean and standard deviation. Significance levels are defined as p < 0.05 (∗), p < 0.01 (∗∗), p < 0.001 (∗∗∗) and p < 0.0001 (∗∗∗∗). Details of the statistical test employed, significance levels, and n numbers are provided in the relevant figure legends.Statistical tests  were calculated using GraphPad Prism versions 7-8; tests used are indicated in Figure Legends. Contingency tables were used when comparing numbers of individual defects e.g., alignment defects in mutants and controls, and P values calculated using Fisher’s exact test (


Structural studies have revealed how GPCRs can activate G proteins. However, whether this mechanism is conserved among all classes of GPCR remains unknown. Here we report the structure of the class-C heterodimeric GABAB receptor, which is activated by the inhibitory transmitter GABA, in its active form complexed with Gi1 protein. We found that a single G protein interacts with the GB2 subunit of the GABAB receptor at a site that mainly involves intracellular loop 2 on the side of the transmembrane domain. This is in contrast to the G protein binding in a central cavity, as has been observed with other classes of GPCR. This binding mode results from the active form of the transmembrane domain of this GABAB receptor being different from that of other GPCRs, as it shows no outside movement of transmembrane helix 6. Our work also provides details of the inter- and intra-subunit changes that link agonist binding to G-protein activation in this heterodimeric complex.G-protein-coupled receptors (GPCRs) have central roles in intercellular communication B receptor in complex with Gi1 protein reveals that the mode of G-protein binding in this class-C G-protein-coupled receptor differs from that of other classes.Cryo-electron microscopy structure of heterodimeric GABA Recent structural studies have provided important information on how GPCRs can act as nucleotide-exchange factors that allow the release of GDP from the inactive G protein, and then the activation of these proteins upon GTP binding2. Several previous structures of activated GPCR–G protein complexes have revealed a similar mode of action for each6. Despite differences in the interaction mode of G proteins for various class-A, -B and -F GPCRs, in all previously characterized interactions the C-terminal extremity of the Gα subunit engages with a cavity on the intracellular side of the receptor that results from the opening of this domain owing to the movement of transmembrane helix (TM) 6 relative to TM37.GPCRs are essential elements that are involved in cell–cell communication and represent major targets for therapeutic drugs8 that are composed of two identical or similar subunits11. These dimers may activate only one G protein at a time11, but the molecular basis of this asymmetric mode of action remains unknown. Among the class-C GPCRs that are activated by the neurotransmitter GABA, the GABAB receptor (hereafter referred to as GABAB) is an attractive drug target for the treatment of brain diseases12. GABAB is composed of two distinct subunits: GB1 (to which agonists bind) and GB2 (which is responsible for G-protein activation)14. Each subunit is composed of an extracellular Venus flytrap (VFT) domain and a transmembrane domain (TMD)15. The structure of this receptor has recently been solved in a number of states, including apo, antagonist-bound, agonist-bound, and agonist- and positive allosteric modulator (PAM)-bound18. Although these studies have helped to identify the conformational changes in subunits that are associated with ligand binding, it remains unclear at the atomic level how this heterodimeric GPCR activates G proteins.Compared to other classes of GPCRs that can be activated in a monomeric form, class-C GPCRs are mandatory dimersB in complex with the G protein Gi1 at 3.5 Å resolution. Our results reveal a mode of G-protein coupling that differs from those that have previously been reported for GPCRs of other classes; our structures reveal that small movements of TM3 and TM5 lead to changes in the intracellular loops (ICLs) that offer a binding site for the G protein on the side of the GB2 subunit of GABAB. These data also help to refine models that describe how agonist binding in the VFT domain of GB1 leads to the activation of the TMD of GB2, and how small molecules can act as PAMs.Here we report the cryo-electron microscopy (cryo-EM) structure of the agonist- and PAM-bound form of the GABA11 19 19 Fig. . Our cryB–Gil complex assumes an overall architecture that is similar to the previously reported low-resolution GABAB–Gil structure in the B2a state11  6UO8) with a root mean squared deviation of 2.3 Å, in which the TMDs adopted a TM6–TM6 interface and the TMD of subunit GB2 showed an outward shift at the intracellular ends of TM3 and TM5. We did not observe conformation changes of TM3 and TM5 in the agonist-bound states (PDB 6UO9)  as a reference, the TMDs of GB1 remained unchanged but the TMDs of GB2 did not overlap well (a root mean squared deviation of 4.8 Å)  and GB2 Y6976.44 had rotamer changes and formed a hydrogen bond with GB2 N6896.45 and GB1 N8116.45, respectively  Fig. . However Å) Fig. . The GB2 Å) Fig. . Therefo3.44 rotated away from TM5 by about 65° to avoid potential spatial clashes   and TM6 (D6886.35) of GB2, all of which are conserved among class-C GPCRs. Similar to class-A GPCRs (in which a D/ERY motif constitutes an ionic lock that stabilizes the inactive state)21, K5743.50 and D6886.35of GB2 form an ionic lock in the inactive state and become weaker owing to inward movement of TM3 upon receptor activation. K5743.50 turned to N5202.39 of GB2 to form an additional ionic interaction  Fig. . Mutatioing Fig. , which sein Fig. . This ision Fig. . Substition Fig. . The int5 helix and the two linker regions (β2–β3 and αN–β1) in Gαi1  and the linker region in β2–β3 (D193) in Gαi1. ICL1 and ICL3 were away from the G protein and participated only in the recognition of the C-terminal ‘hook-like’ region of Gαi  ) by 6–22 fold, which highlights the essential role of ICL2 in GB2–Gi1 coupling (consistent with previous studies22). Substitutions in ICL1 (S515A) or TM3 (R577A and I581W) abolished GABAB-induced production of inositol monophosphate (IP1). Substitutions in TM3  and in ICL3 (L686A) decreased the agonist potency by 9–32 fold, which indicates that ICL1, ICL3 and TM3 are involved in the recognition of Gi1.Given the flexibility of Gax) Fig. . Most muB predominantly couples to Gi/o subtypes of G protein23. The C-terminal 5–9 residues of the α5 helix of G protein have previously been found to be the key determinants for G-protein-coupling specificity25. The α5 helix of Gαi1 contributed 62% (533 Å2) of the interaction surface with GABAB 26  or G352G.H5.24—but not L353G.H5.22 or F354G.H5.25—impaired GABAB-induced G-protein signalling, which suggests an essential role for C351G.H5.23 and a partial involvement of G352G.H5.24 in the specificity of Gi coupling 26 Fig. . We mutaing Fig. , consistGB2 Fig. . When reoss Fig. . The speil binding to GABAB forms a smaller interface (856 Å2) than in the class-A cannabinoid receptor 1 , class-B glucagon receptor (905 Å2), or class-F smoothened receptor  , but rather into a cavity located at the periphery  that comprised the α5 helix, the linker region in β2–β3 and the linker region of αN–β1 28. This binding mode explains the Gi selectivity of the GABAB, and is supported by numerous mutations within the ICLs and the G protein29. The similar determinants that are involved in G-protein recognition, the conservation of ICL231 and the similar mode of activation of these dimeric receptors suggest that there may be a similar coupling mechanism in the other class-C GPCRs.Our cryo-EM structure of the GABAi-bound GABAB structure was almost identical to that observed with the agonist and PAM without Gi  leads to a second movement of the two TMDs with a closer apposition of GB2 on GB1 that creates the PAM-binding site and leads to a change in the bending of the GB2 subunit  signal peptide—including GB1a (UniProt: Q9UBS5) and GB2 (UniProt: O75899)—were cloned into the pEG BacMam vector34. An 8× histidine tag and 3C protease cleavage site were inserted at the C terminus of the GB1a (residues 15–919) subunit, and a Flag epitope tag (DYKDDDD) and a 2× GSG linker were added to the N terminus of the GB2 (residues 42–819) subunit. GABAB and Gil mutants were generated using site-directed mutagenesis. All the constructs were confirmed by sequencing.To facilitate expression and purification, human GABA35. In brief, the 6×histidine-tagged scFv16 was expressed in secreted form in Trichoplusia ni Hi5 insect cells for 48 h using the Bac-to-Bac system. The expressed scFv16 was purified using a Ni-NTA resin. The C-terminal 6×His tag of the Ni-NTA eluent was cleaved by 3C protease and further purified by gel filtration chromatography using a Superdex 200 column. Finally, the purified scFv16 was concentrated and stored at −80 °C until further use.scFv16 was expressed and purified as previously describedi1 was expressed and purified as previously described35. In brief, the dominant-negative Gαi1  and human β1γ2 subunits (β1–8×His tag) were co-expressed in Hi5 insect cells for 48 h using the Bac-to-Bac system. The cells were collected and lysed with a buffer containing 10 mM HEPES (pH 7.5), 100 μM MgCl2 and 10 μM GDP. The cell membrane was collected by centrifugation and heterotrimeric Gi1 was extracted in a buffer containing 1% sodium cholate. The supernatant was purified by Ni-NTA column and the detergent was exchanged with n-dodecyl-β-d-maltoside (Anatrace) on a column. Afterward, Gi1 was mixed with a 1.2 molar excess of scFv16 and further purified by Superdex 200 column. Finally, the Gi1–scFv16 complex was concentrated and flash-frozen in liquid nitrogen until further use.Heterotrimeric G11. The collected cells were solubilized for 3 h at 4 °C in a buffer containing 0.5% (w/v) lauryl maltose neopentyl glycol (Anatrace) and 0.1% (w/v) cholesteryl hemisuccinate (Anatrace). After centrifugation at 30,000g for 30 min, the GABAB was purified by Ni-NTA column and M1 anti-Flag affinity resin. The GABAB was further concentrated and mixed with a 1.3 molar excess of Gi1–scFv16 complex in the presence of 100 μM baclofen and 50 μM BHFF. The sample was incubated at 25 °C for 1 h, followed by the addition of 0.2 U ml−1 apyrase for an additional 1.5-h incubation at 24 °C36. Finally, the sample was purified using a Superose 6 Increase column  to acquire a homogeneous GABAB receptor–Gi1 complex. The entire purification procedure was accomplished in 12 h, followed by immediate verification to acquire a stable and fresh sample for structural determination.The GB1 and GB2 plasmids mixed with PEI 25 K at a 3:0.5:0.5 ratio of PEI to GB1 and GB2 plasmid (w/w) were added to HEK293F cells when the density reached about 2.8 million per ml. Seventeen hours after infection, sodium butyrate was added at a final concentration of 10 mM and the cells were grown for another 3 days at 30 °C before being collectedB–Gi1 complex at 1.8 mg ml−1 was applied onto the glow-discharged holey carbon grids . The grids were blotted for 3.0 s with a blot force of 3 at 4 °C, 100% humidity, and then plunge-frozen in liquid ethane using Vitrobot Mark IV (Thermo Fischer Scientific). Cryo-EM data collection was performed on a Titan Krios at 300 kV accelerating voltage in the Center of Cryo-Electron Microscopy (Zhejiang University). Micrographs were recorded using a Gatan K2 Summit Detector in counting mode with a pixel size of 1.014 Å using SerialEM software37. Image stacks were obtained at a dose rate of about 8.0 electrons per Å2 per second with a defocus ranging from −1.0 to −2.5 μm. The total exposure time was 8 s, and 40 frames were recorded per micrograph. A total of 13,843 movies were collected for the GABAB–Gi1 complex.To prepare cryo-EM grids, 3.0 μl of the purified baclofen- and BHFF-activated GABAB–Gi1 complex were subjected to beam-induced motion correction using MotionCor238. Contrast transfer function parameters for non-dose-weighted micrographs were determined by Gctf39. Cryo-EM data processing was performed using Relion 3.140 and CryoSPARC 2.1541. Template-based particle selection yielded 5,889,932 particle projections using Relion. The projections were imported to CryoSPARC for 2D classification to discard poorly defined particles. The selected particle projections were further subjected to ab initio reconstruction and heterogeneous refinement in CryoSPARC. The well-defined subsets accounting for 1,366,533 particles were re-extracted for further processing in Relion. Three-dimensional classification showed that Gi1 predominantly bound to GB2, however, a small subset  was also found to interact with GB1. To sort out conformational uniform particles for 3D reconstruction, these projections were subjected to 3D classification with a mask on the TMD–Gil, producing one good subset that accounted for 362,826 particles. Further 3D classifications focusing the alignment on the Gil produced two good subsets, which accounted for 275,089 particles that were subsequently subjected to 3D refinement, contrast transfer function refinement and Bayesian polishing. The overall refinement of GABAB–Gi1 generated a map with an indicated global resolution of 3.5 Å at a Fourier shell correlation of 0.143. To further improve the map quality of the complex , local 3D reconstruction focusing on the GABAB receptor and Gi1 was performed using the partial signal subtracted particles in Relion. The local refinement maps for the GABAB and Gi1 showed a global resolution of 3.3 Å and 3.4 Å, respectively, which were combined on the basis of the global refinement map using ‘vop maximum’ command in UCSF Chimera42. This composite map of the GABAB–Gi1 complex was used for subsequent model building and analysis. Global and local resolution was determined using the Bsoft 2.0.7 package43 with half maps as input maps.Image stacks for the GABAB (PDB 7C7Q)11 was used to generate the initial template of the GABAB. The atomic coordinates of Gil and scFv16 from the structure of the human cannabinoid receptor 2–Gi1 complex (PDB 6PT0)36 were used to generate the initial template of the Gi1–scFv16 complex. Models of GABAB and Gil–scFv16 were docked into the electron microscopy density map using UCSF Chimera42. Agonist and PAM coordinates and geometry restraints were generated using a phenix.elbow44. The docked model was subjected to flexible fitting using Rosetta45 and was further rebuilt in Coot45 and real-space-refined in Rosetta45 and Phenix44. The final refinement statistics were validated using the module ‘comprehensive validation (cryo-EM)’ in Phenix. The goodness-of-fit of the model to the map was determined using a global model-versus-map Fourier shell correlation. The refinement statistics are provided in 42 and the UCSF Chimera X package46.The model of the active GABAi1–lgbit, 500 ng Gγ–smbit, 500 ng Gβ, 200 ng GABAB wild type (HA–GB1 and Flag–GB2) or mutants using Lipofectamine 2000 (Thermo Fisher Scientific) in 200 μl of Opti-MEM (Thermo Fisher Scientific). The Flag- and HA-tagged subunits were cotransfected into HEK293T cells and plated in a 96-well plate with a white transparent bottom. HEK293T cells were fixed with 4% paraformaldehyde and blocked with 10% fetal bovine serum (FBS). Bound antibodies coupled to horseradish peroxidase were detected by luminescence using SuperSignal ELISA Femto Maximum Sensitivity substrate (ThermoFisher Scientific), and luminescence was measured using a luminescence microplate reader (Tecan).The cell-surface expression of the receptor subunits was detected using an enzyme-linked immunosorbent assay (ELISA). In brief, HEK293T cells were plated in each well of a 6-well plate at a concentration of 0.3 million per ml (2 ml per well). Plasmid transfection was performed with a mixture of 200 ng GαIP1 accumulation was measured using the IP-One HTRF kit . Transfected HEK293 cells were seeded in a 96-well plate, and 24 h after transfection, cells were treated with baclofen diluted in stimulation buffer in a Cisbio kit for 30 min at 37 °C. Then, cryptate-labelled anti-IP1 monoclonal antibody and d2-labelled IP1 in lysis buffer were added to the wells. After 1 h of incubation at room temperature, the plates were read in PHERAstar FS with excitation at 337 nm and emission at 620 and 665 nm. The accumulation of IP1 was calculated according to a standard dose–response curve.47. The transfection system was the same as that used in the ELISA. After 1 day of transfection, cells in the 6-well plate were digested and resuspended in complete medium DMEM  and plated in 96-well flat-bottomed white microplates. After 24 h, the cells were washed twice with D-PBS and incubated in 40 μl of 5 μM coelenterazine H (Promega) solution diluted with 0.01% BSA- and 5 mM HEPES (pH 7.4)-containing HBSS (assay buffer) for 2 h at room temperature. Baseline luminescence was measured using a luminescent microplate reader (Tecan). The test compound  was added to the cells (10 μl) and incubated for 3–5 min at room temperature before the second measurement. The ligand-induced signal ratio was normalized to the baseline luminescence, and fold-change signals over vehicle treatment were used to show the G-protein dissociation response.G-protein activation was detected using a Nanobit-G protein dissociation assay50), maximum agonist response (Emax) and basal activity for each mutant relative to the wild-type receptor. Data are mean ± s.e.m. from at least three independent experiments, performed in triplicates. ND, not determined. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001  followed by Dunnett’s test, compared with the response of the wild type). For dose–response experiments, data were normalized and analysed using nonlinear curve fitting for the log (agonist) versus response (three parameters) curves.Statistical analyses were performed on at least three individual datasets and analysed using GraphPad Prism software. Bars represent differences in the calculated agonist potency  and Supplementary Table 1.Reporting SummaryVideo 1B and Gi1 represented by the first eigenvector from the principal component analysis.The relative motion of GABAVideo 2B and Gi1 represented by the second eigenvector from the principal component analysis.The relative motion of GABA


The surface roughness of the films varied between 0.6 nm and 4.3 nm, as verified by the Atomic Force Microscopy (AFM) studies. Ammonia (NH3)-sensing measurements of the films were performed for various concentrations at an optimum sensor working temperature of 200 °C. Enrichment of oxygen vacancies confirmed by X-ray Photoelectron Spectroscopy (XPS) in 1% Ag–WO3 enhanced the sensor response from 1.06 to 3.29, approximately 3 times higher than that of undoped WO3. Limit of detection (LOD) up to 500 ppb is achieved for 1% Ag–WO3, substantiating the role of Ag in improving sensor performance.Ag-doped WO Different MOx, such as ZnO, SnO2, WO3, and TiO2 [3) has emerged as a potential MOx to detect the gases, such as NH3, H2, NO2, CO, and alcohol vapors, because of their inherent properties, including excellent electrical conductivity, sensitivity, and selectivity [3-sensing properties of Pd/WO3 films with the response of 0.27 at a working temperature of 225 °C. Lu et al. [2-sensing properties of Ag–WO3 nanoparticles and reported high sensitivity and better selectivity. Xu et al. [3 core-shell nanostructures towards alcohol vapor at a working temperature of 340 °C. All these studies have shown that metal incorporated into WO3 has enhanced the sensor performance.Various types of gas sensors, including chemoreceptive-, ionization-, acoustic-, and resonant-based sensors are gaining more attention in different sectors, such as environmental monitoring, food safety, medical diagnosis, and industrial applications ,2,3. Amoand TiO2 ,5 have bectivity ,7. To enectivity ,9,10. Goectivity  reportedu et al.  presenteu et al.  obtained3-sensing performance of Ag–WO3 films via the spray pyrolysis deposition technique. Ag is a noble metal that enhances the sensor response of WO3 due to electronic sensitization mechanism [3 for NH3 sensing, and one of the works published is on hydrothermal technique [3 films are accessible for NH3 sensing. Therefore, our attempt has proved the possibility of these films for NH3 sensing by spray pyrolysis. Though the literature is available on NH3 sensing by metal-doped WO3, the majority of the sensors work at high temperatures (>250 °C) and the reported detection limit is high [3), a reducing gas, is a major pollutant from the automobiles, fertilizer, and mining industries. According to the OSHA  report, the permissible limit for NH3 is 35 ppm for 15 min [3 films as a promising candidate for NH3 sensing at a working temperature of 200 °C with the detection limit of 500 ppb.In this regard, we have illustrated the NHechanism . Due to echnique . To the  is high ,15,16. Ar 15 min . High ex3) were synthesized by the spray pyrolysis method. For undoped WO3, ammonium metatungstate hydrate (99.99% purity) is dissolved in double-distilled water and a homogeneous solution is obtained. For Ag doping, silver nitrate was taken as a precursor, and doping was done at a 1 wt.%, 3 wt.%, and 5 wt.% ratio. Solution concentration was maintained at 0.01 M. Deposition parameters, such as substrate temperature and flow rate, were kept constant at 400 °C and 1 mL/min, respectively.Undoped and silver-doped tungsten oxide films (Ag–WO4 cm3 by vol.) by purging synthetic air (79% N2 + 21% O2) and NH3 to the sample. The flow of gas was controlled using programmable mass flow controllers (MFCs), and the overall flow was kept constant at 500 sccm. I-V measurements were performed via Keithley source meter 2450 using silver paste electrodes. Sensor response of the films was evaluated using the equation, (a and Rg suggest the film resistance in air and target gas (NH3), respectively. Schematic representation of film synthesis and ammonia-sensing measurements of Ag–WO3 film is represented in the Crystal structure and phase identification were performed via Rigaku SmartLab X-ray diffractometer with Cu Kα radiation at 40 kV, 30 mA. Raman analysis was performed using a Horiba JOBINYVON LabRAM HR spectrometer for the confirmation of structure. Morphological studies were performed via Innova SPM Atomic Force Microscope (AFM). AXIS ULTRA X-ray Photoelectron Spectroscope is used for oxidation state and composition studies. Gas sensing was conducted via dc probe measurements in an enclosed chamber , (202), (140), and (240) planes, indicating monoclinic phase (γ-WO3) of WO3 films (JCPDS card no. 43–1035) [3 films. All the films exhibited ‘a’ axis preferential orientation, i.e., along (200) plane. Evolution of (020) peak centered at 23.4° is observed upon addition of Ag into WO3. The introduction of Ag formed new nucleating centers for WO3, ultimately retaining monoclinic structure. The crystallite size of the films was determined by employing the Scherrer formula [43–1035) ,18. The  formula  and it v formula  were als3 exhibit the same set of XRD peaks, we have conducted Raman measurements for further confirmation [3 films excited with the 532 nm laser source. Clear visibility of peaks at ~112 cm−1, ~134 cm−1, ~270 cm−1, ~325 cm−1, ~719 cm−1, ~806 cm−1, and ~963 cm−1, attributed to the different modes of vibrations in the WO3 lattice. Peaks corresponding to ~719 cm−1 and ~806 cm−1 represent the asymmetric and symmetric stretching vibrational modes (νas and νs) of O–W–O bonds and these are often referred to as the strongest monoclinic WO3 modes [−1 and ~325 cm−1 attribute to the O–W–O bending vibrational modes (δ) and peaks below 200 cm−1 contribute to the lattice vibrational modes [−1, which can be ascribed to the symmetric stretching vibration of terminal W=O bonds possibly associated with the clusters on the film surface [3 films.Since triclinic and monoclinic phases of WOirmation ,21. FiguO3 modes . Peaks lal modes . All the surface ,25. The 3 films examined by AFM in tapping mode configuration. All the samples were scanned in the area of 0.5 × 0.5 µm2. 3. It suggests the coalescence of grains upon Ag incorporation forming larger globules of particles with the presence of voids. At 3% and 5%, Ag doping  were examined via the XPS technique. 7/2 and W 4f5/2 located at the binding energies (Eb) of 35.9 eV and 38.1 eV, respectively, in undoped WO3 film. These represent the +6 oxidation state of W and another satellite peak at 40.5 eV indicates the W 5p3/2 component corresponding to the same oxidation state [3 has led to the formation of +6 and +5 oxidation states of W as depicted in 7/2 and W 4f5/2, respectively, for the +6 oxidation state of tungsten, and other pairs of doublets comprised of W 4f7/2 and W 4f5/2 centered at 32.4 eV and 36.5 eV are comparable to the +5 oxidation state of tungsten [b [3−x.Elemental composition and chemical state of Ag–WOon state . The inctungsten . The peatungsten . If oxyggsten [b ,27. In t1, O2, and O3) in both WO3 and 1% Ag–WO3 films  [The deconvolution of O 1s spectra has produced 3 peaks  ,30. Areao for WO3 and 1% Ag–WO3 was found 35% and 45%, respectively. Hence, the studies inferred that oxygen vacancies increased by 10% in 1% Ag–WO3. The least intense peaks centered at 534.7 eV and 532.8 eV, respectively in WO3 and 1% Ag–WO3, connected to the adsorbed oxygen species  . Figure f silver ,31.3 towards NH3 was fixed by purging gas at various temperatures and calculating subsequent sensor response as presented in 3 is purged at 5 different temperatures: 100 °C, 150 °C, 175 °C 200 °C, and 250 °C. An increase in temperature causes an increase in the sensor current, demonstrating the semiconducting nature of the WO3 films. At 100 °C, the graph shows a straight line, denoting no response for NH3 due to the low thermal energy of the gas required for the chemisorption process. Low response was noted at 150 °C and 175 °C. An increase in the temperature to 200 °C has shown maximum response, and thereafter the response has decreased as indicated in 3 started to respond, as enough thermal energy is provided for the surface reaction to occur by overcoming the activation energy barrier [Operation temperature for WO barrier . After t3 films  for different NH3 concentrations. 3 concentration. The sensor response for 5 ppm NH3 was found to be 1.06, 3.29, 0.34, and 0.62 with the standard error of ±0.07 for undoped, 1%, 3%, and 5% Ag-doped WO3 films, respectively. The highest response was recorded for 1% Ag–WO3 with the 3. ‘Ag’, being a noble metal, acts as a catalyst due to chemical sensitization and also causes electronic sensitization due to the interaction between Ag and WO3, which might have induced the enhancement in the sensor response of the WO3 film [3 showed higher surface roughness value (2-gas sensing, respectively. Augmentation in the sensing performance of 1% Ag–WO3 could be accredited to improved oxygen vacancies upon Ag doping, which is confirmed by XPS analysis and also higher surface roughness and void formation on the film surface is proven by AFM, providing a greater number of adsorption centers for NH3 [2O formation upon Ag doping. When Ag concentration exceeds 1%, Ag2O amounts might increase and cause an increase in the depletion layer width, deteriorating the sensor response of the films.Sensor response, rate of response, and recovery (ss value  compared for NH3 ,35. Neve for NH3 . Also, X3 and 1% Ag–WO3 films towards various gases. These films are tested at a 5-ppm concentration towards NH3, CO, CH4, and NO2. Both films showed the highest response to NH3 indicating the selective nature of the deposited films towards ammonia among other gases. Repeatability measurements of about 5 cycles were performed for both WO3 and 1% Ag–WO3 films, shown in 3 purge at 5 ppm concentration, implying the stable response of the films. In comparison, sensing properties of the current work with the previously reported literature is given in 3 nanoflakes synthesized via spray pyrolysis detected the lowest NH3 concentration to be up to 120 ppm at 150 °C [3 films synthesized by soft chemical route exhibited a detection limit of 100 ppm towards NH3 at 300 °C [3 nanosheets synthesized by acidification with impregnation process detected 2 ppm NH3 at 400 °C [3–Fe2O3 nanocomposites by hydrothermal synthesis demonstrated NH3 detection of 25 ppm at 300 °C [3 gas, keeping an operating temperature of 200 °C, which is a significant improvement in the sensing performance compared to the literature presented in Selectivity and repeatability are the key factors that decide the performance of the sensors. t 150 °C , while tt 300 °C . The Cr–t 400 °C , whereast 300 °C . In the 3 exposure to the synthetic air produces molecular/atomic oxygen ions (3 to the adsorbed oxygen species creates a band bending region called the depletion layer, resulting in a decrease in the carrier concentration of the film and an increase in the resistance. Later, when WO3 is subjected to NH3 exposure, the depletion layer width decreases due to the release of electrons back to the WO3. As Ag–WO3 is exposed to NH3, the depletion layer further decreases due to the electronic sensitization mechanism giving rise to an increment in carrier concentration and a reduction in surface resistance. The well-known gas detection mechanism of metal oxide gas sensors involves the resistance variations caused by the chemisorption of gas molecules on the sensor surface. Depending on the operating temperature, WO3 films were investigated for NH3 sensing at different concentrations. XRD spectra depicted the monoclinic phase of deposited films, which was further verified by Raman analysis. Oxygen vacancies, higher surface roughness, and void-like structures of 1% Ag–WO3 contributed to enhancement in the sensor response value. Selectivity studies of WO3 and 1% Ag–WO3 towards NH3, NO2, CH4, and CO exhibited an excellent response to NH3 compared to other gases. Spray deposited Ag–WO3, as a unique approach for NH3 sensing, produced a superior response at a low operating temperature of 200 °C with a detection limit in the sub-ppm levels.Spray pyrolyzed Ag–WO


S207A,L185D, a first-of-a-kind ‘decoy’ PAR1 inhibitor, by mutating the S207A and L185D residues in wild-type KLK4, which strongly binds to PAR1. KLK4S207A,L185D markedly inhibited PAR1 cleavage, and PAR1-mediated MAPK/ERK activation as well as the migration and invasiveness of melanoma cells. This ‘substrate-capturing’ KLK4 variant, engineered to bind to PAR1, illustrates proof of principle for the utility of a KLK4 ‘proteinase substrate capture’ approach to regulate proteinase-mediated PAR1 signaling.Proteinase-activated receptor-1 (PAR1), triggered by thrombin and other serine proteinases such as tissue kallikrein-4 (KLK4), is a key driver of inflammation, tumor invasiveness and tumor metastasis. The PAR1 transmembrane G-protein-coupled receptor therefore represents an attractive target for therapeutic inhibitors. We thus used a computational design to develop a new PAR1 antagonist, namely, a catalytically inactive human KLK4 that acts as a proteinase substrate-capture reagent, preventing receptor cleavage (and hence activation) by binding to and occluding the extracellular R41-S42 canonical PAR1 proteolytic activation site. On the basis of in silico site-saturation mutagenesis, we then generated KLK4 The activation of PARs is initiated by enzymatic cleavage of their extracellular N-terminal sequence to expose a ‘tethered ligand’ that binds to sites on the extracellular PAR domains to trigger PAR-mediated cell signaling. The first potent activator of PAR1 to be identified and characterized was thrombin, a serine protease that cleaves PAR1 at its ‘canonical’ extracellular cleavage site between residues R41 and S422. It was subsequently found that members of the kallikrein-related peptidase (KLK) family of serine proteinases are also able to cleave/activate PARs3. KLK4, KLK5, KLK6 and KLK13 cleave PAR1 at the canonical R41-S42 site, and KLK13 also cleaves PAR1 at position R46-N474. A variety of other proteinases were found to regulate PAR1 activation by cleaving bonds within the A33 to E57 N-terminal sequence of human PAR1 at distinct sites both upstream and downstream of the ‘tethered ligand’ sequence5. These PAR-regulating proteinases include the matrix metalloproteinases MMP1, MMP2 and MMP13. Of note, the coagulation proteinase, activated protein C (APC), cleaves PAR1 at its R46/N47 site to unmask a tethered ligand sequence, thereby conferring biased signaling that differs from the activation of PAR1 by thrombin6. Thus, the PAR1 N-terminal sequence 33ATNATLDPRSFLLRNPNDKYEPFWE57 constitutes a common target in the sequence of PAR1 for activation by a variety of proteinases.Activation of the G-protein-coupled receptor (GPCR) family of proteinase-activated receptors (PARs) by certain extracellular serine proteinases is known to trigger cell signaling7, inflammation8, and vascular homeostasis9, but it also contributes to some inflammatory10 and cardiovascular pathologies, including atherosclerosis11, restenosis12, and thrombosis13, and to the progression of some cancers by stimulating cell migration and tissue invasion. Indeed, there is a strong correlation between PAR1 expression and the progression of lung14, prostate15, gastric5 and ovarian16 cancers and of melanoma17. In lung, ovarian and breast cancers and in melanoma, the activation of PAR1 promotes invasiveness and tumorigenicity21. For example, Boire et al. showed that MMP1 activation of PAR1 promoted invasiveness and tumorigenesis of breast cancer cells in xenograft models19; Villares et al. showed that in-vivo treatment of melanoma-bearing mice with PAR1 siRNA inhibited melanoma growth and metastasis22; and Adams et al. showed that PAR1 expressed by tumor stromal cells promoted colon cancer growth in vivo23.Following activation by extracellular proteinases, including members of the KLK family, PAR1 plays significant roles in many normal physiological processes, such as coagulation26. Although the KLKs serve as attractive therapeutic targets in the management of certain inflammatory diseases and cancers, the development of selective KLK-targeted enzyme inhibitors represents a challenge27, due to the structural similarity, especially at the catalytic site, of KLK family members. In light of the potential role played by KLK4 (having a sequence homology of 31.64% with thrombin) in prostate cancer via activating either PAR1 or PAR228, we sought to develop a strategy, alternative to generating a KLK4-specific enzyme inhibitor, to prevent KLK4-stimulated activation of PAR1. We thus aimed to prevent the ability of KLK4 to cleave/activate PAR1 by designing a high-affinity catalytically inactive KLK4 that could act as a ‘decoy’ substrate-capturing reagent by binding to PAR1 and thereby masking the activation site and blocking PAR1 activation by the proteolytically active wild-type KLK4.The KLK family of serine proteinases represents the largest cluster of serine proteinases in the human genome. Expressed in a wide variety of tissues, the kallikrein-related peptidases are implicated in numerous pathophysiologies, ranging from cancers to skin disorders, such as Netherton syndrome29 and in inflammatory tissues. Our strategy is derived from the protease ‘substrate-capture or trapping’ approach, as recently reviewed30. This strategy uses catalytically inactive proteinases that bind to, but do not cleave, a substrate. Our approach to inhibitor design comprised engineering an inactive mutant of KLK4 to bind to PAR1 with higher than natural affinity, thus converting it from a PAR1 agonist to a PAR1 antagonist. Since proteases, such as KLK4, are difficult to engineer by applying directed evolution approaches, we utilized a purely computational method for KLK4 engineering before preparing the mutant proteins in the laboratory. Using a computational design, we generated a first-of-a-kind PAR1 inhibitor by mutating the S207A and L185D residues in wild-type KLK4, a natural activator of PAR1 that strongly binds to PAR1. The strong binding of the mutant protein, designated KLK4S207A,L185D, to the extracellular N-terminal region of PAR1 blocked the canonical cleavage site of PAR1, thereby preventing the activation of PAR1 by KLK4. To demonstrate the potential of our newly designed inhibitor of PAR1-activating proteinases present in the tumor microenvironment, we investigated its ability to inhibit PAR1 activation in PAR1-overexpressing cancer cells, and to block the migration and invasion of human melanoma cells lines that endogenously expresses PAR1 17, namely, WM3682 and the highly invasive line WM3314. Our results provide an evaluation of the potential of using the proteinase substrate-capture strategy to block the cleavage/activation of a selected proteinase target, namely, PAR1.Below we describe our novel approach for selectively blocking PAR1 activation by KLK4 and, by extension, other proteinases like the KLKs that might be present in the microenvironment of tumorsThe in-silico saturation mutagenesis approach previously developed in our group allows us to predict affinity-enhancing mutations in any protein–protein interaction by starting from a high-resolution structure of the protein complex. Since the structure of the KLK4/PAR1 complex as such is not available, we modeled the structure of this complex by using the structure of KLK4 and the structure of thrombin (which is homologous to KLK4) interacting with PAR1 (see Methods). Using this modeled structure of the KLK4/PAR1 complex, we identified the binding interface of KLK4, namely, the 14 positions that interact directly with the PAR1 peptide. At these 14 positions, we introduced all possible mutations, one at a time, and computed the change in free energy of binding due to each mutation  and an additional mutation at either position E98 or position L185 . The double KLK4 mutants tested were thus those with the S207A mutation in addition to one of the following mutations: E98I, E98Y, E98F, L185S, L185Q, L185N, L185H, L185R, L185K, L185E, or L185D. The active protease, KLK4WT, and the inactive low-affinity single mutant, KLK4S207A, were also purified and used as controls.The KLK4 variants selected for expression and purification all included the mutation S207A  tagged with a FLAG tag on the N-terminus and a 6 × His tag on the C-terminus. Activation of the enzyme was achieved with thermolysin by cleavage of the bond between the pro-peptide and peptidase domain. Western blot analysis verified the expression of all KLK4 variants except for KLK4S207A,L185S, KLK4S207A,L185N, KLK4S207A,L185R and KLK4S207A,L185K (data not shown), and these mutants were therefore not included in the subsequent experiments. The remainder of the KLK4 protein variants were purified using Ni–NTA affinity column, analyzed by mass spectrometry  analysis of pro-KLK4WT) was determined by measuring the fluorescent signal generated when reacting it with the Boc-VPR-AMC fluorescent substrate  and Koff (3.91 × 10–4 s-1). The addition of an affinity-enhancing mutation, such as L185D , increased the affinity of the soluble protein to the PAR1 peptide by 20-fold (KD = 5.55 × 10–8 M). The affinity enhancement resulted from an improvement in the dissociation rate , despite the slower association rate (Kon = 5.7 × 10 M-1 s-1)  in an in-vitro cellular system, we used the breast cancer MCF7 and melanoma WM3682 cell lines as model systems. The full-length PAR1 protein, fused to an mCherry fluorescent protein at the N-terminus and an eYFP fluorescent protein at the C-terminus, was overexpressed in these cells. As in the wild-type PAR1 protein, the N-terminus (which houses the cleavage site for the KLK4WT peptidase) of the fluorescent fusion PAR1 protein is localized in the extracellular region, and the C-terminus is located in the intracellular region. In both the MCF7 and WM3682 cell lines, cleavage (activation) of PAR1 upon treatment with 100 nM KLK4WT removed the extracellular mCherry, leaving the intracellular eYFP intact , the released mCherry tag appeared to be internalized into the cells. This finding stands in contrast to that for the untreated cells, for which the extracellular domain (mCherry) underwent only a minor cleavage, probably caused by endogenously released extracellular proteases. Notably, addition of 200 nM KLK4S207A,L185D together with 100 nM of the catalytically active KLK4WT inhibited the cleavage of membrane PAR1.To test whether KLK4WT and KLK4S207A,L185D resulted in a significant, dose-dependent inhibition of PAR1 cleavage in comparison with treatment with KLK4WT alone , relative to KLK4WT, were validated using SPR measurements of this mutant in complex with the PAR1 peptide. We thus showed that while a ~ sixfold reduced kon affinity rate constant was observed for the KLKS207,L185D/PAR1 peptide complex compared with that of the KLKS207/PAR1 peptide complex, the koff affinity rate was 123-fold slower for the former, resulting in an 20-fold overall affinity enhancement for the KLKS207,L185D/PAR1 peptide complex.Our findings establish proof of principle for the utility of the proteinase-capture approach to block the cleavage/activation of PAR1, a biologically important KLK4 targeted substrate. Although the proteinase-capture approach has previously been used to identify potential substrates by a ‘pulldown-proteomic’ approach, we believe our work is the first to demonstrate proof of principle for blocking the cleavage/activation of a proteinase substrate, such as PAR1. Using in-silico predictions for affinity-enhancing mutations and experimentally testing several selected double mutants that were based on the catalytically inactive KLK4S207A,L185D ‘decoy’ substrate-capture enzyme was able to block KLK4- mediated cleavage of PAR1 expressed in MCF7 and WM3682 cell lines . Further, treatment of the MCF7 and WM3682 cells overexpressing fluorescently labeled PAR1 with a mixture of KLK4WT and KLK4S207A,L185D resulted in a significant inhibition of PAR1 cleavage in comparison with treatment with KLK4WT alone. Of added importance, inhibition of PAR1 cleavage, presumably caused by endogenously produced microenvironment proteinases, was also observed in the absence of added KLK4WT. This inhibitory effect resulted in reduction in migration induced by unknown PAR1-activating proteinases in the tumor cell microenvironment.The KLK424, was found to be endogenously overexpressed in the MCF7 cell line36. Another member of the MMP family, namely, MMP-13, which activates PAR1 at a different non-canonical cleavage site, between residues S42-F43, was also shown to be expressed in MCF7 cells37. In addition, other members of the KLK family are known to be expressed in MCF7 breast cancer cells38, including: KLK5, which cleaves PAR1 at positions R41-S42 and F43-L44; KLK6, which cleaves PAR1 at positions R41-S42, S42-F43, F43-L44, L45-R46 and R46-N47; and KLK13, which cleaves PAR1 at positions R41-S42, F43-L44, and R46-N473. It is now known that proteinases, such as MMP1, APC, and neutrophil elastase, can cleave PAR1 at ‘non-canonical’ sites to stimulate ‘biased signaling’39. To the best of our knowledge, no previous work has been performed to evaluate protease expression in WM3682 cells, but a screen of the Broad Institute Cancer Cell Line Encyclopedia (CCLE) mRNA expression database revealed that MMP1 is highly expressed in most melanoma cell lines, suggesting that it could also be expressed in the WM3682 cell line and could therefore be involved in the cleavage process of PAR1 at a non-canonical cleavage site, between residues D39 and P4026. The KLK4S207A,L185D ‘decoy’ could, in principle, block PAR1 activation at this site as well as at the ‘canonical’ activation site.Several cell-secreted proteinases have been suggested as enzymes that can cleave/activate PAR1 in MCF7 cells and may thus serve as agonists of PAR1. MMP1, for example, which cleaves PAR1 at a non-canonical site, between residues D39-P40S207A,L185D interacts with the PAR1 extracellular region at the molecular level to block activation at both canonical and non-canonical cleavage sites, we examined the interaction of KLK4S207A,L185D with the PAR1 extracellular region. Structural analysis of the modeled KLK4/PAR1 complex structure showed that the affinity enhancement in the KLKS207A,L185D/PAR1 complex  in complex with KLK4 was modeled on the basis of the available crystal structures of a thrombin/PAR1 complex (PDB-ID 3LU9)42 and the structure of the wild-type KLK4 (PDB-ID 4K8Y)43. Missing residues were added using the Modeller suit44. All water molecules and ions were removed. The crystal structure of KLK4 was aligned with that of thrombin (which is homologous to KLK4) in 3LU9, and the modeled PAR1 peptide was aligned with PAR1 of the same crystal structure. The resulting KLK4/PAR1 complex structure was optimized using FlexPepDock46, and the best ranked model according to the Rosetta generic full-atom energy score was used as the input to the in-silico saturation mutagenesis protocol.The structure of the PAR1 extracellular peptide . For each of the above positions, we defined a set of shell positions that included the residues on KLK4 that are within 3 Å of the KLK4 position to be mutated and the residues on PAR1 that are within 5 Å of the KLK4 position. For each binding interface position, we then performed 17 calculations, in which the considered position on KLK4 was either kept as wild type or was replaced with any other amino acid, except proline, cysteine, or glycine. During the calculation, the shell and the interface residues were repacked, and the energies of the KLK4/PAR1 complexes were calculated for the wild-type and mutated complexes. The two proteins in each complex were then separated to enable calculation of the energy of each unbound chain separately without further repacking of the side chains. The intermolecular energy was calculated by subtracting the energies of the single chains from the total energy of the complex. ΔΔGbind was calculated as the intermolecular energy of the mutated complex minus the intermolecular energy of the wild-type complex. The obtained ΔΔGbind was normalized according to a linear equation obtained in our previous work in which the correlations between various experimental and computed ΔΔGbind values were determined48. Rotamer libraries used for computational modeling were based on the backbone-dependent library of Dunbrack and Karplus49, with additional rotamers expanded by 1 SD around their mean χ1 and χ2 values. For the calculations, we used ORBIT software with the energy function optimized for the design of protein–protein interactions48. The lowest-energy rotameric conformation of each mutated complex was found using the dead-end elimination theorem51.The modeled structure of the KLK4/PAR1 complex was used as the input for all calculations, and hydrogen atoms were added to the structure by MolProbityWT (amino acid sequence and numbering are according to UniProtKB—Q9Y5K2) and selected KLK4 variants  were purified as previously described with minor modifications52. In brief, the pPIC9K plasmid  containing the pro-KLK4WT clone  were isolated from individual Escherichia coli colonies by using a HiYield plasmid mini-kit  and sequenced using primers 1 and 2 (Supplementary data Table WT clone by using site-directed mutagenesis PCR with overlapping primers 3 and 4 (Supplementary Table WT) was digested with the DpnI enzyme, according to the product protocol . The final product (containing only the linear dsDNA product) was electroporated into E. coli, incubated at 37 °C with shaking at 300 rpm for 1 h in LB medium , plated onto LB-Amp plates (ampicillin at 1 mg/L), and then incubated at 37 °C for 24 h. Several colonies were scraped from the plate, transferred into LB-Amp medium, and incubated at 37 °C with shaking at 300 rpm for 24 h. The pPIC9K plasmid containing the KLK4S207A clone was then isolated from each E. coli colony with a HiYield plasmid mini-kit (RBC Bioscience) and sequenced using primers 1 and 2  for chromosomal incorporation by homologous recombination. Transformed cells were plated onto RDB plates  and incubated for three days at 30 °C. Cells were scraped from the plates using E-buffer  and plated onto YPD-G418 plates (4 mg/mL G418) for an additional three days. Colonies (~ 10) from each clone were harvested, seeded into 5 mL of BMGY medium  w/v, 1.1812% KH2(PO4) w/v, 1.34% yeast nitrogen base w/v, 4 × 10–5% biotin w/v, 1% glycerol v/v), and incubated at 30 °C with shaking at 300 rpm for 24 h. For protein expression, cells in BMGY medium were pelleted and then re-suspended in 5 mL of BMMY medium  w/v, 1.1812% KH2(PO4) w/v, 1.34% yeast nitrogen base w/v, 4 × 10–5% biotin w/v, 0.5% MeOH v/v) to reach OD600 of 1.0 and incubated at 30 °C with shaking at 300 rpm for 72 h, with 0.5% MeOH being added every 24 h. Protein expression and secretion into the medium were analyzed by western blot, using mouse anti-FLAG primary antibody  and an alkaline phosphatase conjugated anti-mouse secondary antibody . BCIP reagent  was used for protein expression signal analysis according to the product protocol. Individual clones (representing each of the KLK4 variants) showing the strongest expression levels were selected and subjected to large-scale production and purification procedures. Briefly, clones in 5 mL of BMGY medium at OD600 of 8.0–10.0 were added to 500 mL of BMGY medium at 30 °C with shaking at 300 rpm for 24 h. Cells were precipitated from the medium and induced for protein secretion in 500 mL of BMMY at 30 °C with shaking at 300 rpm for 72 h. MeOH (0.5%) was added to the medium every 24 h. After induction, the supernatant was filtered through a 0.22-µm vacuum filter. The filtrate was then subjected to the following workup: NaCl was added to a final concentration of 300 mM, imidazole  was added to give a final concentration of 10 mM, and the solution pH was adjusted to 8.0. After 1 h at 4 °C, the medium was filtered again, and the protein of interest was purified on a column packed with 5 mL of Ni–NTA Sepharose beads (Invitrogen). The protein was eluted from the Ni–NTA beads with 15 mL of buffer at pH 8.0 containing 50 mM sodium phosphate, 300 mM NaCl and 250 mM imidazole. The eluted protein was concentrated to 5 mL, and the buffer was replaced with PBS using a Vivaspin with a 5-kDa cutoff . The protein was then subjected to activation by thermolysin . Briefly, thermolysin, which cleaves the N'-terminus pro-domain, was added to KLK4 in a mass ratio of 1:80 (thermolysin:KLK4) for 2 h at 37 °C. The reaction was terminated by addition of 1,10-phenanthroline at a final concentration of 10 mM. Activated KLK4 proteins were then purified using gel filtration chromatography on a Superdex™ 75 16/600 GL column in an ÄKTA™ Pure  chromatography system. The gel filtration column was pre-equilibrated with PBS (pH 7.4), and the protein was eluted with PBS. The protein concentration was calculated from protein absorbance at 280 nm  obtained using a NanoDrop spectrophotometer . Non-activated protein samples (not treated with thermolysin), with theoretical mass of 28.6 kDa, were subjected to SDS-PAGE (denatured and reduced) and mass spectrometry analysis . The purified proteins were stored at −80 °C.KLK431SKATNATLDPRSFLLRNPNDKYEPFWED58-FITC, > 95%, Peptron, Daejeon, Korea; residue numbering based on UniProtKB-P25116, PDB-ID 3LU9) was immobilized on the surface of an NLC chip (NeutrAvidin immobilized to a GLC layer) via NeutrAvidin–biotin interactions. The PAR1 peptide (26 µg) dissolved in PBST  was immobilized on the chip to give 2600 response units (RU). An empty channel was used as the negative control. Prior to each binding assay, the temperature was set at 25 °C. Purified KLK4 variants were then allowed to flow over the surface-bound PAR1 peptide at concentrations of 100, 50, 25, 12.5, 6.25 and 0 nM for KLK4WT, concentrations of 2, 1, 0.5, 0.25, 0.125 and 0 µM for KLK4S207A, KLK4S207A,E98I, KLK4S207A,E98Y, KLK4S207A,E98F, KLK4S207A,L185E, KLK4S207A,L185H and KLK4S207A,L185Q, and concentrations of 8, 4, 2, 1, 0.5 and 0 µM for KLK4S207A,L185D, at a flow rate of 30 µL/min for 818 s. The dissociation of the protein-peptide complex was then monitored with PBST flowing over the surface at a flow rate of 30 µL/min for 30 min except for KLK4S207AL185D, where this step was prolonged to 166 min (due to slow dissociation). For each protein complex, a sensorgram was generated from the RUs measured during the course of the interaction minus the values of the empty channel. The values of kon and koff were determined by fitting the sensorgrams to a 1:1 Langmuir kinetic model. The fitted data were considered statistically valid with a χ2 value that was less than the 10% of the RUmax in each of the fitted sensorgrams.The affinity constants for the interaction between the purified KLK4 variants and the PAR1 peptide were determined by SPR spectroscopy on a ProteOn XPR36 (Bio-Rad) instrument at the Cytometry, Proteomic and Microscopy Unit . A biotinylated, PAR1 peptide (biotin-2/dmol) was obtained from the normalized average of four spectra in the range of 200–260 nm. Data points with a diode voltage above 1000 V were excluded.To examine whether any structural change had been caused as a result of the mutations to KLK4, we evaluated the secondary structure of the selected purified non-activated KLK4 proteins by CD spectroscopy . Spectra of 10 μM purified protein in 400 μL of PBS were obtained at 25 °C, and the signal of the blank solvent (PBS) was subtracted. Ellipticity  or the MCF7 human breast cancer cell line . The cells were transfected with a fluorescently labeled PAR1 plasmid (pCDNA3.1 containing the mCherry-PAR1-eYFP construct described previously32) and Lipofectamine™ LTX as a transfection reagent, according to product protocol (Thermo Scientific). For WM3682 cells, following 24 h of transfection, cells were sub-cultured in μ-slide 8-well ibiTreat microscopy chambers  in DMEM containing 10% FBS and incubated for additional 24 h in a humidified incubator at 37 °C and 5% CO2. Cells were washed once with PBS, suspended in FBS-free DMEM, and incubated for another 24 h. Thereafter, the cells were treated with 100 nM KLK4WT (positive control), with 200 nM KLK4S207A,L185D, with a combination of KLK4WT and KLK4S207A,L185D, or with medium alone (negative control) for 1 h under the same conditions as those prior to the analysis. Live fluorescence images were acquired with a confocal laser-scanning microscope  using the Plan Apochromat 63 × /1.4 oil DIC M27 objective . The assay was performed in triplicate, and representative images are presented . For MCF7, the transfection was performed directly on the μ-slide 8-well ibiTreat microscopy chambers (ibidi) and after 24 h, the medium was replaced with FBS-free DMEM and the cells were incubated for another 1 h in a humidified incubator at 37 °C and 5% CO2. Thereafter, the cells were treated with 100 nM KLK4WT, 200 nM KLK4S207A,L185D, 200 nM KLK4S207A, a combination of KLK4WT and KLK4S207A,L185D or KLK4WT and KLK4S207A, or with medium alone (negative control) for 1 h and incubated under the same conditions as those prior to the analysis. Cells were washed once with PBS, then fixed in 4% paraformaldehyde in PBS for 10 min at room temperature, washed again, and covered with PBS. Fluorescent samples were imaged using an FV1000 laser scanning confocal microscope (Olympus) with 488 and 561 nm solid state lasers and a × 40/1.25 NA silicon immersion objective . Fluorescence images were acquired using the Flowview 10 ASW V2.1 software. The assay was performed in two replications (separate wells for each condition), and two fields from each well were examined.To follow the cleavage of full-length cell-expressed PAR1 by KLK432 and incubated for 24 h at 37 2 in room air. Transfected cells grown to confluency in the two T25 flasks were then lifted with 0.25% trypsin in 1 mM-EDTA-supplemented PBS, pH 7.4. Cells were combined and subcultured in a 48-well multi-dish tray (Nunc 1 cm2 area) in 10% serum-supplemented DMEM and incubated for a further 24 h. Cells were serum starved by replacing the culture medium with 0.2 mL of serum-free DMEM containing 0.1% bovine serum albumin (BSA) and incubated overnight. The next day, cells were washed 3 times with isotonic Hanks’ balanced salt solution pH 7.4 containing 10 mM HEPES, 1.5 mM MgCl2, 1.5 mM CaCl2, and 0.1% BSA and replenished with 0.1 mL of the same buffer. KLK4WT and KLK4S207A,L185D were mixed and preincubated for 1 h on ice at 10 times the final specified concentration added to the cells. In each well, 10 µL of pre-incubated 10 × enzyme mix was then added to the indicator cell monolayer containing 90 µL of buffer and incubated for 30 min at 37 S207A,L185D treated cells was determined by a one-way ANOVA for measurements performed for four cell monolayer replicates.Two T25 flasks of confluent WM3682 cells were transfected with the mCherry-PAR1-YFP construct described previously6 cells were seeded in a 6-well plate in DMEM containing 10% FBS and incubated in a humidified incubator at 37 °C and 5% CO2 until reaching 80% confluence. The cells were then washed with a flow-cytometry buffer (0.1% BSA in PBS), harvested using a cell scraper, and incubated for 30 min in a flow-cytometry buffer or in a flow-cytometry buffer containing anti PAR1 Alexa Fluor 680 conjugated antibody . Cells were washed once with a flow-cytometry buffer and analyzed using a flow cytometer. For the negative control, the same procedure was carried out with one exception: 0.25% trypsin solution was used to detach the cells from the 6-well plate (instead of a cell scraper). Analysis was carried out on an Accuri C6 Flow Cytometer , and data analysis was performed using a FlowJo software .The expression levels of PAR1 in WM3682 and WM3314 cells were determined using flow cytometry. In brief, 0.3 × 10S207A,L185D of the migratory phenotype of the WM3682 cell line was performed by using a scratch assay, as previously described53. Briefly, WM3682 cells (1.5 × 105) were cultured as confluent monolayers in a 24-well plate in DMEM containing 10% FBS and incubated in a humidified incubator at 37 °C and 5% CO2. When the cells reached 100% confluence, a scratch was formed by removing a strip of cells across the well with a p200 pipette tip. The scratched monolayers were then washed twice with PBS to remove non-adherent cells, and 500 µL of DMEM supplemented with 2% FBS, containing 12.5, 25, or 200 nM KLK4S207A,L185D or PBS (control), was added. The wells were captured with an EVOS FL cell imaging system at × 4 magnification, both immediately after cell wounding (t0) and again after 24 h (t24). The experiment was performed in triplicate, and the images were analyzed using ImageJ software. The ratio between the gap areas at t0 and t24 was calculated.Testing the inhibition by KLK453. Briefly, ThinCert 24-well cell culture inserts  were coated with growth factor reduced Matrigel  diluted in DMEM  at a 1:30 v/v ratio. The lower chamber was filled with 600 μL of DMEM supplemented with 10% FBS . WM3314 cells , in the presence or absence of KLK4S207A,L185D protein, were incubated in 200 µL of FBS-free DMEM, added to the pre-coated ThinCert cell culture inserts, and incubated for 18 h at 37 °C and 5% CO2. Invasive cells were stained with a DippKwik stain kit (American MasterTech Scientific) and detected by an EVOS FL Cell Imaging System , at ×4 magnification. The experiment was performed in triplicate, and the images were taken from the center field of each insert. Analysis of digitized images was performed using ImageJ software and a Cell Colony Edge Analyser.An in-vitro Boyden chamber assay was performed as previously described with minor changes54. The cells were grown in complete DMEM  containing 10% FBS, penicillin, streptomycin, and l-glutamine until they reached 80% confluence. Cells were scraped with a cell scraper, diluted 1:2, and grown overnight in complete DMEM. Thereafter, 2.5 × 105 cells were transferred into a 12-well plate with complete DMEM for 24 h. At this point, the medium was replaced with a starvation medium (FBS-free complete DMEM) for 18 h. After starvation, the cells were washed with PBS and incubated in a starvation medium containing 10 nM KLK4WT and 0, 10, 100 and 1000 nM KLK4S207A,L185D in a humidified incubator at 37 °C and 5% CO2 for 15 min. The positive control contained 10 nM KLK4WT alone, and the negative control was incubated in starvation medium without any added protein. The cells were transferred to ice, and a lysis buffer  was added. The cells were detached with a cell scraper, collected, incubated on ice for 10 min, and centrifuged at 14,000 g for 30 min, and the supernatants were transferred to fresh tubes. Western blot analysis was performed for all samples using anti-MAP kinase (Erk1/2) produced in rabbit or anti-MAP kinase, activated (diphosphorylated Erk1/2) antibody produced in mouse as a primary antibody . A secondary HRP-linked anti-rabbit or anti-mouse antibody was then added, and the signal was developed using the EZ-ECL kit . The chemiluminescent signal was imaged with Fusion FX . Equal amounts of total protein from cell lysate samples were loaded in each lane on the SDS-PAGE, as measured by a colorimetric BCA protein assay kit . The experiment was performed in triplicate, and a representative membrane image was selected for presentation.The experiment was performed using the WM3682 cell line as previously described with minor modificationsS207A,L185D on the growth and survival of WM3682 and MCF7 cells were assessed by an XTT assay . WM3682 or MCF7 cells were seeded (7500 cells per well) in a 96-well microplate  and incubated at 37 °C at 5% CO2 for 24 h. The medium was then replaced with fresh DMEM supplemented with 10% FBS, penicillin, streptomycin, and l-glutamine, and the cells were incubated for 24 h at 37 °C at 5% CO2 in the presence or absence of 1 μM KLK4S207A,L185D. Viable cells from each condition were measured by reading the absorbance at 450 nm (UV), using a plate reader, as described in the manufacturer’s protocol. The UV readings of the cell-only control were normalized to 100%, and readings from cells treated with KLK4S207A,L185D were presented as a percentage of the control.The effects of KLK4p value < 0.05 was considered statistically significant.Data were analyzed with GraphPad Prism version 5.00 for Windows . Data shown in all the figures are the means of triplicates from independent experiments, and error bars represent the standard error of the mean. Statistical significance was determined by column statistics and one-way ANOVA analysis. A Supplementary Information.