PPFIA1 binds to and controls active α5β1 integrin recycling

We have previously identified Nrp1 and additional adaptor and motor proteins as key components of a signalling complex that selectively promotes endocytosis of active α5β1 integrin and the polymerization of FN into a basolateral fibrillar network by ECs11,14. While these findings highlight endocytic trafficking events involving active α5β1 integrins11,14 for the turnover of FN fibrils9,10, it is unclear whether the endocytosed active α5β1 integrins only remove fibrillar FN or whether they also replenish it. To address this question, we determined the subcellular localization of the ubiquitously expressed adaptor protein PPFIA1 that has been shown to couple endocytosis and exocytosis in presynaptic nerve termini32. Standard confocal microscopy analysis revealed that, as previously reported33, fluorescently immunostained PPFIA1 localized around mature peripheral focal adhesions of ECs (Fig. 1a, magnified panel 1). Furthermore, PPFIA1 also localized to the vicinity of centrally located vinculin-containing fibrillar adhesions (Fig. 1a, magnified panel 2), where active α5β1 integrins, as recognized by SNAKA51 monoclonal antibody (mAb)34,35, predominantly localized14. Similarly, when transfected in ECs, PPFIA1-GFP concentrated in close proximity to focal (Fig. 1b, magnified panel 1) and fibrillar (Fig. 1b, magnified panel 2) adhesions. These findings were further substantiated by super-resolved timed-gated stimulated emission depletion (g-STED) confocal microscopy analyses of PPFIA1-GFP transfected and anti-vinculin-stained or anti-active α5β1 integrin-stained ECs (Fig. 1a,b, g-STED).

Figure 1: PPFIA1 localizes in proximity of EC fibrillar adhesions and selectively interacts with active α5β1 integrin cytodomain. (a,b) Conventional or g-STED (boxed) confocal microscopy analyses of endogenous PPFIA1 (green) or PPFIA1-GFP (green) and vinculin (red) or SNAKA51+ active α5β1 integrin (red) in ECs indicates that PPFIA1 localizes close to both focal and fibrillar adhesions. Right-sided panels 1 (focal adhesions) and 2 (fibrillar adhesions) are magnifications of boxed areas in left-sided panels. (c) Representation of inactive and active α5β1 integrin displaying clasped or unclasped cytotails, respectively. Epitopes recognized by VC5 and SNAKA51 mAbs are shown. VC5 binds to a β propeller domain epitope of the α5 subunit of both inactive and active α5β1 integrin, while SNAKA51 binds to the calf domains of the α5 subunit only when α5β1 integrin is active. Jun or Fos N-terminally-tagged recombinant integrin cytotails were exploited to mimic α5β1 integrin activation state. Jun-Fos dimerization and clasping of integrin cytotails, mimicked inactive α5β1 integrin. (d) Immunoprecipitation from EC lysates of total (that is, inactive and active) or active α5β1 integrin by VC5 and SNAKA51 mAbs, respectively, followed by western blotting with anti-PPFIA1 or anti-α5 integrin subunit or anti-Nrp1 antibodies. In ECs, PPFIA1 associates with active, but not inactive α5β1 integrin in both siCTL and siNRP1 ECs. Total EC lysates (input) were employed for control purposes. (e) Pull-down from mouse fibroblast cell lysates of PPFIA1 on isolated or dimerized α5 or β1 integrin cytotails. Kindlin-2 was pulled down for control purposes. Both PPFIA1 and kindlin-2 are pulled down on isolated Myc-Fos β1, but not Flag-Jun α5 integrin subunit cytotail. Furthermore, both interactions are severely impaired by forced Jun-Fos-mediated dimerization of α5 and β1 integrin subunit cytotails. α5 or β1 integrin cytotails are revealed by means of Western Blot with the respective anti-tag antibodies (Flag and/or Myc) or anti-β1 integrin subunit cytotail (for loading control purposes). Scale bar, 20 μm (a,b) and 2 μm (boxed g-STED panels). Full size image

To test whether PPFIA1 represents a novel regulator of active α5β1 integrin recycling, we first assessed whether PPFIA1 binds α5β1 integrins, and if NRP1 mediates this interaction. Lysates from ECs oligofected with either control siRNA (siCTL) or a pool of four siRNA targeting NRP1 (siNRP1) were immunoprecipitated with mAbs recognizing either total (that is, active and inactive, clone VC5) or active (clone SNAKA51) α5β1 integrins36 (Fig. 1c, left panel). PPFIA1 co-immunoprecipitated with SNAKA51+/active α5β1 integrins, but NRP1 did not mediate the interaction (Fig. 1d). Indeed, quantitative confocal analysis of relative surface fluorescence of individual ECs showed that SNAKA51+ active α5 integrin subunits account for the 10% of VC5+ or AB1928+ total α5 integrin subunits (Supplementary Fig. 1).

To corroborate the interaction between active α5β1 integrin and PPFIA1, we performed pull-down assays from cell lysates using recombinantly expressed and purified monomeric or dimeric α5 or β1 integrin cytotails. Complementary DNAs encoding the cytoplasmic domain of either α5 or β1 integrin subunit were fused at their N-terminus with Jun and Fos peptides, respectively (Fig. 1c, right panel). To mimic the unclasped and active or the clasped and inactive α5β1 integrin conformation, recombinant α5 and β1 cytotails were employed alone or in combination to force their dimerization and clasping via the high affinity Jun-Fos interaction (Fig. 1c, right panel). The well-known β1 integrin cytosolic interactor kindlin 2 (ref. 37) was used as a positive control. Both PPFIA1 and kindlin 2 were pulled down by isolated Fos β1, but not Jun α5 integrin subunit cytotails. Furthermore, the interactions of both proteins were significantly impaired by forced Jun-Fos-mediated dimerization of α5 and β1 integrin subunit cytotails (Fig. 1e).

In ECs, PPFIA1 localizes in close proximity to fibrillar adhesions and interacts with the β1 cytotail of active α5β1 integrins. Hence, we sought to investigate whether, in line with the role it plays in neuronal synapses31,32, PPFIA1 participates in the recycling of endocytosed active α5β1 integrins. By means of biochemical assays38, we first analyzed the extent of total or active α5β1 integrin recycling at different time points on internalization. In accordance with previous quantitative fluorescence confocal microscopy analyses39, we found that total α5β1 integrin rapidly recycled at early (5 min), but not late (10–15 min) time points after internalization (Supplementary Fig. 2a). By contrast, active α5β1 integrin was more efficiently recycled back to the EC surface at late (15 min) rather than early (5 min) time points after internalization (Supplementary Fig. 2b). Next, we studied the effects of PPFIA1 silencing (siPPFIA1, Fig. 2a) on active α5β1 integrin recycling in ECs. pLVX lentivirus-driven overexpression of an siRNA-resistant PPFIA1 (pLVX-PPFIA1r) was employed to rescue PPFIA1 expression levels (Fig. 2a). Notably, when compared with control transduced (pLVX) and silenced (siCTL) ECs, PPFIA1 knockdown (pLVX siPPFIA1) significantly reduced the amount of recycled active α5β1 integrin (Fig. 2b). Importantly, pLVX-PPFIA1r restored the defective recycling phenotype of PPFIA1-silenced (pLVX-PPFIA1r siPPFIA1) ECs (Fig. 2b).

Figure 2: PPFIA1 drives the recycling of endocytosed active α5β1 integrin. (a) Western blot analysis of EC lysates, transduced with pLVX lentivirus carrying or not the silencing-resistant PPFIA1 construct (PPFIA1r) and then control (siCTL) or PPFIA1 (siPPFIA1) silenced. (b) Time-course analysis of the relative amounts of recycled active α5β1 integrin in pLVX siCTL ECs versus pLVX siPPFIA1 ECs or siPPFIA1 ECs rescued with pLVX-PPFIA1r, evaluated by integrin recycling assay and capture ELISA assay. PPFIA1 silencing significantly impairs the recycling of active α5β1 integrin. Values are mean±s.e.m., n=3 independent experiments (two technical replicates for each experiment). (c) Confocal microscopy analysis of anti-active α5β1 integrin mAb SNAKA51 localization (red) in living confluent ECs following 20 min of incubation. SNAKA51+ active α5β1 integrin localizes in fibrillar adhesion in pLVX siCTL, but not in pLVX siPPFIA1 ECs in which it accumulates instead in perinuclear punctae. PLVX-mediated PPFIA1r overexpression restores the fibrillar adhesion localization of SNAKA51 in siPPFIA1 ECs. The number of active α5β1 integrin-containing adhesions per 100 μm2 of cell area was quantified in pLVX siCTL, pLVX siPPFIA1 or pLVX-PPFIA1r+siPPFIA1 ECs. Data are mean values±s.e.m., n=20 cells per condition pooled from two independent experiments. (d) Fluorescent confocal microscopy analysis of EC perinuclear punctae in which SNAKA51+ active α5β1 integrin accumulates on PPFIA1 silencing. In siPPFIA1 ECs, SNAKA51+ active α5β1 integrin is enriched in a perinuclear vesicular compartment labelled by both TGN (TGN46) and early endosome (EEA1), but not late endosome (LAMP1) markers. Relative amount of fluorescence intensity of SNAKA51+ active α5β1 integrin, respectively on EEA1+ endosomes or on TGN46+ PGCs, was evaluated. Data are mean value±s.e.m., n=70 cells per condition pooled from two independent experiments **P<0.01; ***P<0.001; Student’s t-test. Scale bar, 50 μm (c), 10 μm (d). Full size image

To determine whether impaired active α5β1 integrin recycling may correlate with its altered subcellular localization, we incubated SNAKA51 mAb for 20 min at 37 °C on confluent living pLVX siCTL or pLVX siPPFIA1 ECs that were then processed for immunofluorescence and analyzed by confocal microscopy. In comparison with pLVX siCTL ECs, where SNAKA51 mainly localized in α5β1-containing fibrillar adhesions, in pLVX siPPFIA1 ECs SNAKA51+/active α5β1 integrins accumulated in perinuclear punctae (Fig. 2c). Reintroduction of PPFIA1r rescued the fibrillar distribution of active α5β1 integrin in PPFIA1-silenced (pLVX-PPFIA1r siPPFIA1) ECs (Fig. 2c). Furthermore, fluorescence confocal microscopy analyses of perinuclear punctae revealed that, in siPPFIA1 ECs, internalized active α5β1 integrin accumulated in TGN46+ vesicles, which correspond to bona fide post-Golgi carriers (PGCs) and, in accordance with previous findings40,41, are often positive for the early endosome antigen 1 (EEA1) (Fig. 2d, upper panels). The negligible fraction of internalized SNAKA51+/active α5β1 integrins that colocalize with the late endosome marker lysosomal-associated membrane protein 1 (LAMP1) was not affected by PPFIA1 silencing (Fig. 2d, lower panels).

Unlike non-polarized fibroblasts42, FN fibrils are deposited beneath the basolateral surface of polarized ECs17,18,19. Similarly to what has been previously reported for active αvβ3 integrin after shear stress43, quantitative analysis of apico-basal mean intensity ratio by confocal xz sectioning on both sparse (Supplementary Fig. 3) and confluent ECs (Fig. 3a–c) revealed that SNAKA51+/active α5β1 integrins localize along the basolateral side of pLVX siCTL ECs, while in pLVX siPPFIA1 ECs they were randomly redistributed around the cell surface (Fig. 3a–c). Also in this case, lentiviral transduction of PPFIA1r rescued the defective polarized membrane distribution of SNAKA51+/active α5β1 integrin caused by PPFIA1 silencing in ECs (Fig. 3a–c). Furthermore, confocal xz sectioning demonstrated that also PPFIA1-GFP resides along the basolateral side of confluent ECs (Supplementary Fig. 4).

Figure 3: PPFIA1 drives basolateral localization of active α5β1 integrin. (a) Confocal xz sectioning microscopy analysis of anti-active α5β1 integrin cell surface localization (red) following 20 min of incubation with SNAKA51 mAb on living confluent ECs. (b) Schematic representation of the quantification of apico-basal mean intensity ratio of SNAKA51+ active α5β1 integrin distribution on the EC surface. b.l., basolateral. (c) Quantitative analysis of apico-basal mean intensity ratio of SNAKA51+ active α5β1 integrin. SNAKA51+ active α5β1 integrin localizes on the basolateral surface of pLVX siCTL ECs, but not pLVX siPPFIA1 ECs in which it randomly redistributes all around the cell surface. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. Scale bar, 5 μm (a) ***P<0.001; Student’s t-test. Full size image

Taken together, these data indicate that in ECs PPFIA1 selectively associates with and regulates the trafficking of SNAKA51+/active α5β1 integrins.

PPFIA1 controls FN secretion and polymerization

Since α5β1 integrin is the principal FN receptor in ECs and its localization in fibrillar adhesions is fundamental to funnel actomyosin tension necessary to unfold and incorporate secreted FN dimers into polymeric fibrils8, we asked whether the polymerization of extra domain-A (ED-A)-containing cellular FN (ED-A FN) was controlled by PPFIA1. Confluent pLVX siCTL, pLVX siPPFIA1 and pLVX-PPFIA1r siPPFIA1 ECs were cultured in medium containing plasma FN-depleted fetal bovine serum (FBS) and endogenous ED-A FN was visualized with the IST9 mAb and confocal fluorescence microscopy. While pLVX siCTL ECs polymerized ED-A FN into an extracellular fibrillar network (Fig. 4a, upper panels), which confocal xz sectioning localized to the basolateral membranes (Fig. 4b), pLVX siPPFIA1 ECs accumulated FN in a perinuclear compartment inside ECs (Fig. 4a, middle panels). Importantly, pLVX-mediated PPFIA1r overexpression strongly decreased ED-A FN intracellular accumulation and restored its polymerization (Fig. 4a, lower panels). Addition of exogenous FN to the medium did not rescue the defective fibrillogenesis of endogenous ED-A FN in PPFIA1-silenced ECs (Supplementary Fig. 5).

Figure 4: PPFIA1 controls endogenous ED-A FN exit from the TGN and its polymerization underneath confluent ECs. (a) Confocal microscopy analysis of IST9 mAb-labelled endogenous ED-A FN (green) in confluent ECs. ED-A FN polymerizes into a fibrillar network in pLVX siCTL, but not in pLVX siPPFIA1 ECs in which it accumulates in perinuclear punctae. PLVX-mediated PPFIA1r overexpression restores ED-A FN polymerization in siPPFIA1 ECs. The relative amount of fibrillar ED-A FN area was calculated in pLVX siCTL, pLVX siPPFIA1 and pLVX-PPFIA1r+siPPFIA1 ECs. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. (b) Confocal xz sections of ED-A FN (green) in siCTL or siPPFIA1 ECs. Compared with siCTL ECs, the basolateral accumulation of polymerized ED-A FN decreases dramatically in siPPFIA1 ECs. (c) Confocal microscopy characterization of the perinuclear compartment in which ED-A FN accumulates on PPFIA1 silencing in ECs. Before fixation, living ECs were incubated for 20 min with exogenous SNAKA51. Compared with siCTL ECs, ED-A FN (blue) heavily accumulates in the TGN46+ (green) TGN cisternae of siPPFIA1 ECs. Of note, in siPPFIA1 ECs exogenously added SNAKA51 mAb (red) binds active α5β1 integrins that, on endocytosis, reach TGN46+ post-Golgi carrier vesicles, but do not enter in TGN cisternae. (d) Fluorescent confocal microscopy shows that, similarly to SNAKA51, exogenously added rhodamine (Rhod, red) labelled serum FN reaches TGN46+ (green) post-Golgi carrier vesicles, but does not enter in TGN cisternae of siPPFIA1 ECs in which endogenous ED-A FN (blue) heavily accumulates instead. (e) Western blot analysis of soluble ED-A FN released by confluent ECs seeded on Transwell inserts. An equal percentage of apical and basolateral volumes of medium were collected after 72 h of culture from different wells of siCTL or siPPFIA1 ECs. Equal amounts of exogenous rabbit IgG were added to samples (spike normalization) for loading control purposes. Quantification of the ratio between apical or basolateral amount of ED-A FN released by siCTL over siPPFIA1 ECs. PPFIA1 silencing impairs basolateral, but not apical ED-A FN secretion. Data are mean±s.e.m., n=8 wells per condition pooled from 4 independent experiments. Scale bar, 50 μm (a), 10 μm (c,d), 5 μm (b). ***P<0.001; Student’s t-test. Full size image

Next, we investigated whether the subcellular compartment in which ED-A FN accumulated on PPFIA1 silencing overlapped with the compartment(s) in which active α5β1 integrins accumulated in siPPFIA1 ECs. To this end, living cells were incubated with SNAKA51 mAb at 37 °C for 20 min, fixed, stained and analyzed by fluorescence confocal microscopy. Interestingly, in contrast to pLVX siCTL ECs, pLVX siPPFIA1 ECs displayed a large accumulation of ED-A FN in TGN46+ TGN cisternae. Furthermore, ED-A FN staining overlapped with internalized SNAKA51 mAb-bound, active α5β1 integrin only in TGN46+ vesicular PGCs, but not in TGN cisternae of pLVX siPPFIA1 ECs (Fig. 4c). Moreover, exogenously added rhodamine-labelled plasma FN reached the TGN46+ positive PGCs of siPPFIA1 ECs, but did not enter the TGN cisternae where instead endogenous ED-A FN accumulated (Fig. 4d). Next, we explored if PPFIA1 silencing affects not only the assembly of ED-A FN into a basolateral fibrillar meshwork, but also its polarized secretion. To this end, we set up a quantitative biochemical apico-basal secretion assay that, through the use of Transwell polycarbonate membrane inserts, allowed quantification of the amount of soluble ED-A FN that confluent ECs released in either the apical or basolateral culture medium. In keeping with previous findings17,18,19, we observed that the secretion of endogenous ED-A FN mainly occurred at the basolateral side of ECs (Fig. 4e). Notably, PPFIA1 silencing severely impaired basolateral, but not apical, ED-A FN secretion in the medium of confluent cultured ECs (Fig. 4e).

Altogether, these data show how in ECs the lack of PPFIA1 simultaneously impairs the recycling of endocytosed active α5β1 integrin and the basolateral secretion of endogenous ED-A FN. Furthermore, PPFIA1 silencing results in an abnormal accumulation of both ED-A FN and endocytosed active α5β1 integrin in PGCs. With respect to the role of PPFIA1 in the docking of secretory vesicles to the plasma membrane of neuronal presynaptic terminals31,32, we asked whether TGN46+ PGCs might also reach the basolateral plasma membrane of ECs and localize in close proximity to adhesion sites where PPFIA1 concentrates (Fig. 1; ref. 33). Confocal microscopy analysis on fixed ECs revealed a sizeable accumulation of TGN46+ PGC vesicles around vinculin-containing adhesion sites where PPFIA1 also localizes (Fig. 5a). To evaluate further the possible targeting of PGCs in close proximity (100 nm) to adhesion sites, we performed time-lapse total internal reflection fluorescence (TIRF) microscopy on living ECs co-transfected with the GFP-tagged TGN marker sialyltransferase (ST-GFP, gift of Julia von Blume) and the Cherry-tagged adhesion site component vinculin (Cherry-vinculin, gift of Kenneth Yamada). ST-GFP localized in typical perinuclear TGN cisternae (Fig. 5b, white arrowhead), as well as in more peripheral vesicular PGCs, several of which were observed targeting close to Cherry-vinculin-containing adhesion sites (Supplementary Movie 1; Fig. 5b; Supplementary Fig. 6a). Similarly, TIRF microscopy revealed that ST-GFP+ PGCs are also delivered in close proximity to PPFIA1-Cherry-positive structures (Supplementary Movie 2; Fig. 5c; Supplementary Fig. 6b) that concentrate around vinculin-CFP-containing ECM adhesions (Fig. 1; ref. 33). Quantitative analyses of snapshots from live time-lapse TIRF microscopy unveiled in siCTL, but not in siPPFIA1 ECs, a statistically significant preferential targeting of ST-GFP+ PGCs in proximity of Cherry-vinculin-labelled ECM adhesions, as opposed to Cherry-vinculin-devoid non-adhesive plasma membrane regions (Fig. 5d).

Figure 5: Post-Golgi carrier vesicles target EC adhesion sites. (a) Confocal microscopy analysis on fixed ECs shows that, in addition to labelling the perinuclear TGN cisternae, TGN46 (green) is present in post-Golgi carrier vesicles that preferentially accumulate, together with PPFIA1 (red), around vinculin-containing (blue) adhesion sites. (b) Snapshots from live TIRF microscopy of a region of an EC that was co-transfected with the TGN marker ST-GFP (green) and the adhesion site component Cherry-vinculin (red). ST-GFP is clearly localized at the TGN cisternae (white arrowhead) and in peripheral post-Golgi vesicles, the trajectory of one of which (white circle) is tracked from the perinuclear area to a peripheral Cherry-vinculin-labelled adhesion site. Single channel photograms are shown in Supplementary Fig. 3. (c) Snapshots from live TIRF microscopy of a region of an EC that was co-transfected with ST-GFP (green), PPFIA1-Cherry (red) and the vinculin-CFP (blue). The trajectory of one ST-GFP post-Golgi vesicles (white circle) is tracked in proximity of a CFP-vinculin labelled adhesion site (white arrow), where it reaches a PPFIA1-Cherry+ area (white arrowhead) and resides for at least 12 s. More in general, several ST-GFP-labelled vesicles closely associate to PPFIA1 surrounded vinculin-containing adhesion sites (Supplementary Movies 1 and 2). Single channel photograms are shown in Supplementary Fig. 3. (b,c) Lower panels are magnifications of the white circled areas shown in the above panels. White arrows indicate representative ST-GFP+ PGCs that are reaching PPFIA1-Cherry+ areas. (d) Quantification of the relative amount of ST-GFP vesicles residing in focal adhesion containing (FA) or non-containing (non-FA) areas in siCTL or siPPFIA1 ECs. Data are mean±s.e.m., n=10 cells per condition pooled from three independent experiments. Scale bar, 10 μm (a–c). ***P<0.001; Student’s t-test. Full size image

To sum up, when PPFIA1 is absent, endocytosed active α5β1 integrins accumulate in PGCs and integrin recycling to the EC surface slows down. Furthermore, on PPFIA1 silencing, a significant fraction of endogenous ED-A FN arrests in the TGN and fails to polymerize. Thus, PPFIA1 controls the recycling of internalized active α5β1 integrin as well as the polarized basolateral secretion and polymerization of newly synthesized endogenous ED-A FN in ECs.

Fibrillar adhesions and FN fibrils are dynamic structures

Since impairing the polarized basolateral secretion of newly synthesized ED-A FN diminishes FN fibrillogenesis (Fig. 4a), we hypothesized that the presence of the FN network underneath confluent ECs depends on its continuous turnover, which in turn hinges on a fine balance between endocytosis-mediated removal of old and cleaved FN fragments and exocytosis-mediated deposition of new FN dimers. To explore this hypothesis directly, we sought to impair the endocytosis of active α5β1 integrins from fibrillar adhesions and determine how this affects FN fibril formation in PPFIA1-silenced ECs. To interfere with active α5β1 integrin internalization, we focused on the small GTPase Rab21, which has been reported to control active β1 integrin endocytosis44,45 and localizes both on early endosomes and at the TGN46. We found Rab21 to control the endocytosis of endogenous ED-A FN in ECs (Fig. 6a).

Figure 6: Fibrillar adhesions and FN fibrils are dynamic structures governed by Rab21 and PPFIA1. (a) Living siCTL and siRAB21 ECs were incubated with IST9 mAb for 30 min, fixed, acid washed and stained. RAB21 silencing impairs ED-A FN endocytosis as revealed by the decrease of IST9 punctae co-localizing with the endosome marker EEA1. Right panels are magnifications of the boxed areas in left panels. Data are mean value±s.e.m., n=70 cells per condition pooled from two independent experiments. ***P<0.001. (b) Left panel, western blot analysis of PPFIA1, RAB21 and actin on total lysates of siCTL, siPPFIA1 and siPPFIA1+siRAB21 ECs. Right panel, western blot analysis of the insoluble matrix fraction of ECs that were extracted with DOC buffer. PPFIA1 silencing dramatically reduces the amount of DOC-insoluble fraction of endogenous ED-A FN. Of note, simultaneous silencing of Rab21 GTPase (siPPFIA1+siRAB21), which drives integrin endocytosis, rescues the defective incorporation of endogenous ED-A FN in the DOC-insoluble fraction of siPPFIA1 ECs. (c) Confocal microscopy analysis of the patterning of endogenous cellular ED-A FN (green) in fixed confluent ECs. Before fixation, living ECs were incubated for 20 min with exogenous SNAKA51 (red). ED-A FN polymerizes into a fibrillar network in siCTL, but not in siPPFIA1 ECs. Simultaneous silencing of Rab21 GTPase (siPPFIA1+siRAB21) fully restores ED-A FN polymerization in siPPFIA1 ECs. SNAKA51+ active α5β1 integrin localizes in fibrillar adhesion in siCTL, but not in siPPFIA1 ECs. Notably, simultaneous Rab21 (siPPFIA1+siRAB21) silencing promotes the localization of SNAKA51 in fibrillar adhesion of siPPFIA1 ECs. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. Scale bar, 20 μm (a,c, right), 50μm (c, left). ***P<0.001; Student’s t-test. Full size image

Western blots of deoxycholate (DOC) buffer-extracted insoluble ED-A FN from confluent siCTL, siPPFIA1 and siPPFIA1+siRAB21 ECs revealed that PPFIA1 silencing dramatically reduced the amount of DOC-insoluble ED-A FN in ECs, while simultaneous silencing of PPFIA1 and Rab21 (siPPFIA1+siRAB21) rescued the defective incorporation of endogenous ED-A FN in the DOC-insoluble fraction of siPPFIA1 ECs (Fig. 6b). Confocal microscopy showed that, compared with siCTL ECs (Fig. 6c, top left panels), ED-A FN failed to polymerize into a fibrillar network in siPPFIA1 ECs (Fig. 6c, middle left panels), while simultaneous silencing of Rab21 GTPase (siPPFIA1+siRAB21) fully restored ED-A FN polymerization in siPPFIA1 ECs (Fig. 6c, bottom left panels). Moreover, fluorescence confocal microscopy analysis revealed that, after incubation on confluent living ECs for 20 min at 37 °C, SNAKA51+/active α5β1 integrin localized to fibrillar adhesion in siCTL (Fig. 6c, top right panels) , but not in siPPFIA1 ECs (Fig. 6c, middle right panels). Importantly, simultaneous PPFIA1 and Rab21 (siPPFIA1+siRAB21) silencing reestablished the localization of SNAKA51 in fibrillar adhesions of siPPFIA1 ECs (Fig. 6c, bottom right panels).

Thus, the plasticity of fibrillar adhesions and the deposition of FN fibrils depend on the functional connection of Rab21 and PPFIA1 that respectively signal to control the endocytosis and recycling/exocytosis of active α5β1 integrin and cellular ED-A FN.

PI4KB/AP-1A/PTPRF-driven FN release and active α5β1 traffic

To identify proteins involved in the recycling of endocytosed active α5β1 integrin in ECs, we investigated the role of crucial regulators of PGC biogenesis21, such as phosphatidylinositol 4-kinase, catalytic, beta (PI4KB) and the EC-expressed clathrin adaptor protein complex-1A (AP-1A). In addition, we characterized the function of PTPRF that directly binds28 and cooperates with PPFIA1 in the docking of TGN-derived neurotransmitter vesicles to the active zone of neuronal presynaptic nerve terminals31,32. To this end, we examined by confocal microscopy the deposits of endogenous cellular ED-A FN in confluent ECs silenced with siCTL or a pool of four siRNA directed against either PTPRF (siPTPRF) or PI4KB (siPI4KB) (Supplementary Fig. 7). Endogenous ED-A FN polymerized into a fibrillar network in siCTL (Fig. 7a, top left panels), but neither in siPTPRF (Fig. 7a, middle left panels) nor in siPI4KB (Fig. 7a, lower left panels) ECs. Similarly, SNAKA51+/active α5β1 integrin localized in fibrillar adhesion of siCTL, but not in siPTPRF or siPI4KB ECs (Fig. 7a, right panels). Quantitative analysis of apico-basal mean intensity ratio of SNAKA51+/active α5β1 integrin by confocal xz sectioning on confluent ECs revealed that SNAKA51+/active α5β1 integrins localized on the basolateral surface of siCTL. However, in siPPFIA1, siPTPRF, or siPI4KB ECs SNAKA51+/active α5β1 integrins redistributed randomly all around the cell surface (Fig. 7b). For control purposes, we silenced the liprin β protein PPFIBP1, that similarly to PPFIA1 binds PTPRF and control synaptic size30, but whose function in organizing the traffic of presynaptic nerve terminals is poorly understood29. We found that PPFIBP1 silencing only modestly affected the apico-basal distribution of SNAKA51+/active α5β1 integrin in ECs (Fig. 7b). Furthermore, silencing of either PI4KB (Fig. 7c) or the key AP-1A subunit AP1M1 (ref. 47) (siAP1M1) (Fig. 7d) or PTPRF (Fig. 7e) significantly impaired the recycling of SNAKA51+/active α5β1 integrin. As in the case of active α5β1 integrin apico-basal distribution, knocking down PPFIBP1 affected the recycling of SNAKA51+/active α5β1 integrin much less efficiently (Fig. 7f).

Figure 7: PTPRF cooperates with PI4KB and AP-1A to control ED-A FN exit from the TGN and polymerization as well as active α5β1 integrin recycling. (a) Confocal microscopy analysis of the patterning of endogenous cellular ED-A FN (green) in fixed confluent ECs and anti-active α5β1 integrin mAb SNAKA51 (red) incubated on living confluent ECs for 20 min before fixation. ED-A FN polymerizes into a fibrillar network in siCTL, but neither in siPI4KB nor in siPTPRF ECs. SNAKA51+ active α5β1 integrin localizes in fibrillary adhesion in siCTL, but neither in siPI4KB nor in siPTPRF ECs. Relative amount of fibrillary ED-A FN area was measured in siCTL, siPTPRF and siPI4KB ECs. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. The number of active α5β1 integrin-containing adhesions per 100 μm2 of cell area was quantified in siCTL, siPTPRF and siPI4KB ECs. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. (b) Quantitative analysis of apico-basal mean intensity ratio of SNAKA51+ active α5β1 integrin by confocal xz sectioning on confluent ECs. SNAKA51+ active α5β1 integrin localizes on the basolateral surface of siCTL, but neither siPPFIA1, nor siPTPRF, nor siPI4KB ECs in which it randomly redistributes all around the cell surface. Silencing of PPFIBP1 affects the apico-basal distribution of SNAKA51+ active α5β1 integrin much less efficiently. (c–f) Time-course analysis of the relative amounts of recycled active α5β1 integrin in siCTL ECs versus siPPFIBP1 or siPTPRF or siAP1M1 or siPI4KB ECs, as evaluated by integrin recycling assay and capture ELISA assay. Silencing of either PTPRF or PI4KB or AP1M1 significantly impairs the recycling of SNAKA51+ active α5β1 integrin. Data are mean±s.e.m., n=3 independent experiments (two technical replicates for each experiment). Scale bar, 50 μm (a, left), 20 μm (a, right), 5 μm (b). *P<0.05; **P<0.01; ***P<0.001; Student’s t-test. Full size image

Altogether, these data indicate that, similarly to PPFIA1 (Fig. 4), PI4KB, AP-1A and PTPRF control the recycling of active α5β1 integrin as well as the polymerization of a FN fibrillar network underneath cultured ECs. Furthermore, the data show that the internalized active α5β1 integrins exploit the same PI4KB/AP-1A-dependent trafficking pathway for recycling.

Rab11B drives FN secretion and active α5β1 recycling

The results above indicated how in ECs, the same molecular and cellular mechanisms control active α5β1 integrin recycling as well as the basolateral secretion and polymerization of ED-A FN. Rab11 small GTPases, which localize at recycling endosomes, TGN and PGCs48, regulate the slow recycling of active β1 integrins13,39. Next, we investigated if in ECs Rab11 proteins may simultaneously orchestrate active α5β1 integrin recycling together with ED-A FN secretion and fibrillogenesis. We found that ECs express both Rab11A and Rab11B proteins (Supplementary Fig. 7). In polarized epithelial cells, Rab11A resides in a vesicular compartment distinct from that of Rab11B49 and, Rab11A, but not Rab11B, controls the apical delivery of post-Golgi cargoes50. When compared with siCTL ECs, Rab11B (siRab11B), but not Rab11A (siRab11A) knockdown halved the amount of recycled active α5β1 integrin in ECs (Fig. 8a). Similarly, on incubation on living confluent ECs, SNAKA51 mAb largely localized in α5β1-containing fibrillar adhesions of siCTL and siRab11A, but not siRab11B ECs (Fig. 8b). In addition, quantitative confocal xz sectioning analysis on confluent ECs revealed that although SNAKA51+/active α5β1 integrins localize along the basolateral side of siCTL and siRab11A ECs, they redistributed randomly around the surface of siRab11B ECs (Fig. 8c). In addition, while siCTL and siRab11A ECs polymerized ED-A FN into an extracellular fibrillar network, siRab11B ECs accumulated FN in a perinuclear compartment, consisting of TGN46+ TGN cisternae (Fig. 8d). Moreover, in siRab11B ECs SNAKA51+/active α5β1 integrins accumulated in perinuclear TGN46+ PGCs (Fig. 8d). Finally, quantitative biochemical apico-basal secretion assays revealed that Rab11B silencing preferentially impaired basolateral ED-A FN secretion in the medium of confluent cultured ECs (Fig. 8e). We hence infer that in ECs the Rab11B small GTPase orchestrates the recycling of endocytosed active α5β1 integrins together with the basolaterally-polarized secretion and polymerization of endogenous ED-A FN.

Figure 8: RAB11B controls the recycling of endocytosed active α5β1 integrins and polarized ED-A FN secretion in ECs. (a) Time-course analysis of recycled active α5β1 integrin in siCTL ECs versus siRAB11A ECs or siRAB11B ECs. RAB11B, but not RAB11A, silencing significantly impairs active α5β1 integrin recycling. Data are mean±s.e.m., n=3 independent experiments (two technical replicates for each experiment). (b) Confocal microscopy analysis of anti-active α5β1 integrin mAb SNAKA51 localization (red) in living confluent ECs (20 min of incubation). SNAKA51+ active α5β1 integrin localizes in fibrillar adhesion in siCTL and in siRAB11A, but not in siRAB11B ECs in which it accumulates in perinuclear punctae. The number of active α5β1 integrin-containing adhesions per 100 μm2 of cell area was quantified in siCTL, siRAB11A and siRAB11B ECs. Data are mean values±s.e.m., n=20 cells per condition pooled from two independent experiments. (c) Confocal xz sectioning microscopy analysis of anti-active α5β1 integrin mAb SNAKA51 localization (red) in living confluent ECs (20 min of incubation). Quantitative analysis of apico-basal mean intensity ratio reveals that SNAKA51+ active α5β1 integrin localizes on the basolateral surface of siCTL and siRAB11A, but not of siRAB11B ECs, where it redistributes all around the cell surface. Data are mean values±s.e.m., n=20 cells per condition pooled from two independent experiments. (d) Confocal microscopy analysis of IST9 mAb+ endogenous cellular ED-A FN (green) in confluent ECs. ED-A FN polymerizes into a fibrillar network in siCTL and siRAB11A, but not in siRAB11B ECs, where it accumulates in a perinuclear compartment. Relative amount of fibrillary ED-A FN area was calculated in siCTL, siRAB11A and siRAB11B ECs. Data are mean values±s.e.m., n=20 cells per condition pooled from 2 independent experiments. (e) Western blot analysis of soluble ED-A FN released by confluent ECs seeded on Transwell inserts. An equal percentage of apical and basolateral volumes of medium were collected after 72 h of culture from different wells of siCTL or siRAB11B ECs. Equal amounts of rabbit IgG were exogenously added to samples (spike normalization) for loading control purposes. Quantification of the ratio between apical or basolateral amount of ED-A FN released by siCTL over siRAB11B ECs. RAB11B silencing much more severely impairs basolateral than apical ED-A FN secretion. Data are mean±s.e.m., n=6 wells per condition pooled from three independent experiments. Scale bar, 50 μm (d), 20 μm (b), 5μm (c). **P<0.01; ***P<0.001; Student’s t-test. Full size image

α5β1 controls FN secretion and polymerization

Altogether, the above results led us to postulate that α5β1 integrin may play a key direct role in the control of ED-A FN secretion. To verify this hypothesis, we investigated whether interfering with α5β1 integrin function in ECs may indeed hamper the secretion of ED-A FN from the TGN and its assembly into a basolateral fibrillar network. Therefore, to hinder directly α5β1 integrin function, we silenced the α5 integrin subunit (siITGA5) in ECs (Fig. 9a). Of note, in addition to the expected severe impairment of FN fibrillogenesis, the silencing of α5 integrin subunit also promoted a clear accumulation of endogenous ED-A FN in the TGN of ECs (Fig. 9b). Furthermore, quantitative biochemical apico-basal secretion assays unveiled that α5 integrin subunit silencing significantly decreases the basolateral, but not apical secretion of ED-A FN in the medium of confluent cultured ECs (Fig. 9c). Thus, blocking α5β1 integrin function in ECs inhibits the basolateral secretion of endogenous ED-A FN from the TGN.

Figure 9: α5β1 regulates ED-A FN secretion and polymerization. (a) Western blot analysis of lysates of ECs control (siCTL) and α5 integrin subunit (siITGA5) silenced ECs. Cells were lysed 24 hours after the second siRNA oligofection and proteins were separated by SDS–PAGE and probed for α5 integrin subunit or actin (for control purposes). (b) Confocal microscopy analysis of IST9 mAb-labelled endogenous ED-A FN (green) in confluent ECs. ED-A FN polymerizes into a fibrillar network in siCTL, but not in siITGA5 ECs in which it accumulates in the TGN46+ (red) TGN cisternae. The relative amount of fibrillar ED-A FN area was calculated in siCTL and siITGA5 ECs. Data are mean±s.e.m., n=20 cells per condition pooled from two independent experiments. ***P<0.001; Student’s t-test. (c) Western blot analysis of soluble ED-A FN released by confluent ECs seeded on Transwell inserts. An equal percentage of apical and basolateral volumes of medium were collected after 72 h of culture, from different wells of siCTL or siITGA5 ECs. Equal amounts of exogenous rabbit IgG were added to samples (spike normalization) for loading control purposes. Quantification of the ratio between apical or basolateral amount of ED-A FN released by siCTL over siITGA5 ECs. α5 integrin subunit silencing impairs basolateral, but not apical ED-A FN secretion. Data are mean±s.e.m., n=8 wells per condition pooled from four independent experiments. **P<0.01; Student’s t-test. Scale bar, 50 μm (b). Full size image

PPFIA1 drives vascular morphogenesis in vitro and in vivo

Polarized FN secretion and matrix polymerization are required for vascular morphogenesis in vitro7 and for vessels formation in vivo in the developing mouse embryo1. First, we directly tested the influence of PPFIA1 on vascular morphogenesis in vitro by means of EC capillary formation assays in growth factor-reduced Matrigel. After 8 h incubation at 37 °C, pLVX siCTL ECs formed capillary networks surrounded by a dense meshwork of polymerized cellular ED-A FN. By contrast, pLVX siPPFIA1 ECs failed to form a fibrillar FN network and remained immobile on the Matrigel surface without forming cell-to-cell contacts and a capillary network. Importantly, pLVX-mediated PPFIA1r re-expression rescued FN network formation, EC mobility and the formation of capillary networks (Supplementary Fig. 8).

Next, we assessed the effects of PPFIA1 inactivation in vivo. Antisense morpholino oligonucleotides (MOs) were used to knockdown Ppfia1 protein expression in the transgenic zebrafish line Tg(kdrl:EGFP) expressing EGFP in ECs. To this end, a translation blocking MOs (experimental group; MO-ppfia1) and the corresponding five-base mismatch MOs (control group; MO-CTL) were independently microinjected into fertilized zebrafish eggs at the single-cell stage. At 48 h post-injection (hpi), the gross appearance of ppfia1 morphant embryos was normal. However, approximately 35% of them exhibited cardiovascular defects, such as an enlarged heart chamber phenotype due to atrium dilation associated with pericardial oedema, blood stasis (Fig. 10a), and reduced blood flow despite the presence of cardiac activity (Supplementary Movies 3 and 4). In contrast, control-injected embryos appeared phenotypically normal. Furthermore, sporadic haemorrhages in the subintestinal vascular plexus, malformations in the intersegmental vessels (Se) and irregular shapes (margin) of the dorsal aorta (DA) and the posterior cardinal vein (PCV) were also present in defective ppfia1 morphants (Fig. 10a). At 72 hpi, all these phenotypic cardiovascular alterations persisted, but did not further exacerbate.

Figure 10: PPFIA1 silencing affects vascular morphogenesis in developing zebrafish embryo. (a) Lateral view at conventional light and confocal fluorescence microscopy of Tg(kdrl:EGFP) zebrafish embryos carrying EC-specific EGFP expression and derived from fertilized zebrafish eggs that were injected with 83 μM of a ppfia1 translation blocking morpholino (MO-ppfia1), 5-base mismatch (MO-CTL), or ppfia1 translation blocking morpholino and 40 pg PPFIA1 mRNA at the single-cell stage. Starting from 48 hpi, ∼35% of the ppfia1 morphants displayed: (i) an enlarged heart chamber phenotype (red arrows) due to atrium dilation associated with pericardial oedema, blood stasis and reduced blood flow despite the presence of cardiac activity; (ii) malformations in the intersegmental vessels (Se, yellow arrow); (iii) an irregular shape (margin) in the DA and the PCV (white arrows). Human PPFIA1 mRNA co-injection successfully rescued the cardiovascular defects in the vast majority (∼84%) of ppfia1 morphants that appeared morphologically normal (b). Western blot analysis of MO-ppfia1, MO-CTL and MO-ppfia1+PPFIA1 mRNA morphants, at 48 hpi, revealed that the band corresponding to zebrafish ppfia1 present in MO-CTL, completely disappear in MO-ppfia1 and it is partially restored in MO-ppfia1+PPFIA1 mRNA morphants. Actin was used as loading control. (c) Relative percentage of the embryos having normal or altered phenotype in two independent experiments is reported in the graph and both absolute numbers with relative percentage of the embryos for each experimental group are reported in Supplementary Tables 1 and 2. (d) Confocal fluorescence microscopy analysis of MO-CTL, MO-ppfia1 and MO-ppfia1+ PPFIA1 mRNA Tg(Kdrl:EGFP) zebrafish embryos trunk cross-sections at 72 hpi stained with DAPI and Fn1a. Right panels are magnifications of boxed areas of the left panels. While MO-CTL morphants show normal Fn1a expression around the PCV, DA, gut and in the whole trunk cross-section (upper panels); MO-ppfia1 morphants display a decreased Fn1a expression in the overall trunk region (middle panels). Co-injection of human PPFIA1 mRNA in MO-ppfia1 morphants rescued Fn1a deposition defects albeit not completely in the overall trunk cross-section. Relative amount of Fn1a expression in MO-CTL, MO-ppfia1 and MO-ppfia1+ PPFIA1 mRNA Tg(Kdrl:EGFP) zebrafish whole trunk cross-sections or around PVC and DA, at 72 hpi. Data are mean values±s.d. from 4 (MO-CTL) or 9 (MO-ppfia1 and MO-ppfia1+ PPFIA1 mRNA) embryos from two independent experiments, and 5th/95th percentile.*P<0.05; **P<0.01; ***P<0.001; ANOVA followed by the Tukey range test. Scale bar, 100 μm (d, left panels), 25 μm (d, right panels). Full size image

To exclude the possibility that cardiovascular aberrations of ppfia1 morphants might be due to MO off-target effects, we assessed whether co-injecting human PPFIA1 mRNA, which was not targeted by either ppfia1 MOs, could rescue the observed phenotypes51. We analyzed wild type embryos, ppfia1 morphants and ppfia1 morphants co-injected with human PPFIA1 mRNA, confirming its effective translation into the corresponding PPFIA1 protein by western blot, although at lower levels than that of endogenous Ppfia1 in wild type embryos (Fig. 10b). As previously observed, about 35% of 48 hpi ppfia1 morphants exhibited the described cardiovascular defects, while control-injected embryos were unaffected. Human PPFIA1 mRNA co-injection successfully rescued the cardiovascular defects in the large majority (∼84%) of ppfia1 morphants that appeared morphologically normal (Fig. 10a,c), while a minor fraction of them (∼16%) still exhibited cardiovascular abnormalities.

Next, we directly evaluated whether the lack of Ppfia1 may affect the deposition of FN in the living zebrafish. To this end, we performed quantitative fluorescence confocal microscopy analyses of FN-immunostained trunk sections of 72 hpi wild type embryos, ppfia1 morphants and ppfia1 morphants co-injected with human PPFIA1 mRNA. As shown in Fig. 10d, ppfia1 morphants exhibited a sizeable reduction in FN deposition both around DA and PCV and in overall trunk sections (Fig. 10d). Co-injection of human PPFIA1 mRNA in ppfia1 morphants rescued FN deposition defects albeit not completely (Fig. 10d), suggesting a dose-dependent threshold for the different phenotypic alterations observed in ppfia1 morphants (Fig. 10b).

In summary, PPFIA1 depletion inhibits endothelial FN secretion and polymerization, and impairs vascular morphogenesis in cultured ECs and in the developing zebrafish embryo.