Integrins are required for stress-fibre attachment and control of F-actin levels

Because loss of integrins causes lethality, we used mitotic recombination to create cells lacking βPS, and examined stress-fibre distribution in clones of follicle cells derived from these mutant cells. As focal adhesions precede the formation of stress fibres in cells in culture (Couchman and Rees, 1979), we expected the integrin adhesive junctions to be required for the formation of stress fibres in follicle cells. We were therefore surprised to find that, in clones of cells lacking βPS, prominent bundles of F-actin were still present at the basal surface (Fig. 4A-B′). These bundles were not organized into the normal parallel arrangement and were displaced towards the periphery. The amount of F-actin at the basal surface of the mutant cells was greater than in adjacent wild-type cells, with quantification of the images revealing a 33% increase (Fig. 4G). Thus, integrins are not required to generate basal actin fibres, but instead are needed to reduce fibre number and organize them into parallel arrays. In addition to the change in stress fibres, lack of βPS-containing integrins resulted in changes to the shape of the follicle cells. The cells lost their hexagonal shape (Fig. 4D), and were unable to flatten fully, as revealed by the position of the nuclei in a lower focal plane (Fig. 4C,C′) and their shape in section (Fig. 4E). The morphology of the mutant cells within the columnar epithelium at earlier stages seemed normal (Fig. 4F) (Devenport and Brown, 2004). In some cases, integrin loss was accompanied by extensive filopodia-like actin spikes (Fig. 5A′).

Open in a separate window

Fig. 4.

Integrin adhesion is required for stress-fibre organization and the control of F-actin content at the basal surface. (A-F) Micrographs of clones lacking βPS, with clones marked either by the absence of GFP (A,B; no green) or the presence of GFP (C-F; green). (A′,B′) Show actin distribution; (A,B) show the merge of actin (red) and the position of the clone. (A,A′) Stage 6 egg chamber showing mutant cells forming thicker filaments than wild-type neighbours. (B,B′) Basal F-actin is stronger in cells lacking βPS at stage 12. (C,C′) In the absence of integrin, basal F-actin fails to assemble into stress fibres (C) and cells fail to flatten as indicated by nuclei at a lower focal plane (C′). (D) A mid-cell section shows the rounded shape of the cells lacking integrin (actin, red; nuclei, blue); the yellow lines show the shape of wild-type cells. (E,F) Transverse sections of follicular epithelia [actin, red; nuclei, blue; CD8-GFP, green (mutant cells); actin channel, grey). Red bars on the right indicate the thickness of the epithelia and apical (`A') and basal (`B') surfaces. The oocyte membrane is just above `egg'. Green asterisks on the actin channel indicate the position of mutant cell(s). Cells lacking integrin were rounded and failed to flatten at stage 12 (E), whereas stage 10A cells seemed normal (F). (G) Quantification of basal F-actin in cells lacking integrins (βPS) compared with wild type (WT) measured as integrated area (grey mean value multiplied by cell area). Wild-type value (92) divided by mutant value (122) is 0.75, which indicates a 33% increase in basal actin in the mutant cells. P=2.8×10^–5 (Student's t-test two samples assuming equal variance). Scale bars: 20 μm. Scale bar shown in A is for A,A′; scale bar shown in B is for all other micrographs except D.

Open in a separate window

Fig. 5.

Integrins downregulate actin-nucleating factors. Micrographs show clones lacking βPS, marked by the presence of GFP (green) in stage 12 egg chambers, except for C-C′″, which is stage 10B. (A-A′″) In addition to not assembling F-actin into fibres, cells lacking integrins form F-actin aggregations associated with Enabled (blue arrow). (B-B″) Mid-cell section showing that cytoplasmic profilin is increased in cells lacking integrins (B″), and F-actin distribution reveals rounded cell shape (B′) [cytoplasmic profilin was increased throughout the cell (data not shown)]. (C-C″) Diaphanous levels were increased at the basal surface of cells lacking integrins. Scale bars: 20 μm.

We next explored possible mechanisms to explain the increase in F-actin in cells lacking integrins, by examining the distribution of proteins that stimulate actin-filament assembly: Enabled (Ena), profilin and Diaphanous (Dia) (Chakraborty et al., 1995; Reinhard et al., 1995; Butler et al., 2006). Ena was previously shown to localize at adhesion sites and genetic removal of Ena, profilin or Dia reduces basal F-actin in follicle cells (Baum and Perrimon, 2001) (data not shown). Furthermore, Ena overexpression produces aggregates of F-actin and Ena in both mammalian and follicle cells (Gertler et al., 1996; Baum and Perrimon, 2001), and similar aggregates were observed in cells lacking integrins (Fig. 5A-A″). Ena accumulated within these aggregates at levels higher than within focal adhesions in adjacent wild-type cells (Fig. 5A″), suggesting elevation of Ena levels. Cytoplasmic profilin was increased in mutant cells, so that the higher concentration of nuclear profilin seen in wild-type cells was no longer apparent (Fig. 5B-B″). This regulation occurs post-transcriptionally, because profilin mRNA levels were unchanged (data not shown). Basal surface levels of Dia were also increased, with this elevation clearer at earlier stages (Fig. 5C-C″). Similar to profilin, this occurred post-transcriptionally, because dia mRNA levels were unchanged (data not shown). Assay of monomeric actin levels by DNAse-I staining showed no difference between wild-type and mutant cells (data not shown), suggesting that the F-actin increase is not driven by increasing monomer concentration. Thus, integrin function is essential for regulating the amount of F-actin at the basal surface and its organization into ordered parallel arrays. This is correlated with an increase in levels of the three actin-polymerization factors Ena, Dia and profilin, suggesting that integrins normally downregulate the actin machinery to control the amount of F-actin incorporated into stress fibres.

αPS1βPS and αPS2βPS have distinct roles

Having characterized the role of all βPS-containing integrins in the organization of follicle-cell stress fibres, we then examined the role of the two major integrin heterodimers expressed in follicle cells, αPS1βPS and αPS2βPS [expression of αPS3 or αPS4 mRNA was not detected in most follicular cells, whereas αPS5 mRNA was only detected at late stages 11 and 12 (Dinkins et al., 2008)]. We first examined the mechanism that results in the transition from one integrin to the other, and found that this occurs by mRNA regulation. The mRNA encoding αPS1 was first detected in follicle cells from stage 4, was retained through stage 11 and then decreased (Fig. 6A). The mRNA encoding αPS2 showed reciprocal expression, becoming first detectable at stage 10B and remaining from then onward (Fig. 6B). Expression of αPS2 mRNA at an earlier stage using the Gal4 system resulted in early production of αPS2βPS (data not shown), demonstrating the lack of a post-transcriptional mechanism to prevent early production of αPS2βPS. A similar switch of integrins in Caenorhabditis elegans is regulated by the transcription factor Pax6 (vab-3) (Meighan and Schwarzbauer, 2007). However, testing whether this is conserved was thwarted by possible redundancy between the four Drosophila Pax6 orthologues, all of which are expressed in follicle cells (supplementary material Fig. S1).

Open in a separate window

Fig. 6.

αPS1βPS and αPS2βPS have distinct functions for follicle-cell stress-fibre organization. (A,B) In situ hybridization on whole egg chambers showing in black the levels of mRNAs encoding αPS1 (A) and αPS2 (B) during oogenesis. (C-D′) Clones of cells lacking αPS1, marked by the absence of GFP. (C,D) Position of the mutant cells (not green) are shown together with the distribution of actin (red) and another protein if relevant (blue). (C,C′) Cells lacking αPS1 form fibres with brighter actin intensity than in wild-type cells; compare arrowheads. (D,D′) Loss of αPS1 elevates αPS2 levels; compare arrowheads. (E-E″) Overexpression of αPS2 using the UAS-GAL4 system increased αPS2 levels (E″) but did not change the actin fibres (E′). (F-G′) Clones of cells lacking αPS2, marked by the presence of GFP (MARCM clones of if^B4). (F,F′) Loss of αPS2 does not perturb stress fibres; compare arrowheads pointing to a wild-type and a mutant cell on F′. (G-G′) αPS1 level in focal adhesions was increased when αPS2 was removed; compare arrowheads in G′. (H) Quantification of basal F-actin in cells lacking αPS1 (mew) compared to wild type (WT) as shown in C and measured as integrated area (grey mean value multiplied by the cell area). Wild-type value (106) divided by mutant value (147) is 0.72, which indicates a rough increase of 38% in basal actin in the mutant cells, comparable to the increase observed in the total absence of integrins (see Fig. 2E). P=1.5.E-8 (Student's t-test two samples assuming equal variances). (I-I′″) Re-expression of αPS1 (blue, I′″) in cells lacking αPS2 (CD8-GFP, green, I″) and stained for actin (red, I′) and αPS2 (green also, I″) results in a delay in stress-fibre re-orientation at stage 12 (I′); most stress fibres in CD8-GFP-positive cells have failed to reorient compared to WT neighbours. (J-J″,L) Global (J) and magnified (J′) view of the basal surface of an egg chamber at early stage 12 where the whole follicular epithelium lacks αPS2 and re-expresses αPS1. In this extreme case, the stress fibres have random orientation yet the whole egg morphology is normal (J″) compared to a wild-type egg chamber (K). Scale bar in B: 50 μm (for A,B,J″,K); scale bar in F: 20 μm (for C-I′″); scale bars in J and J′: 20 μm.

Next, we removed each integrin heterodimer, by making clones of cells mutant for each α-subunit, and examined the consequences at late stages. In both, stress fibres were still organized, but cells that lacked αPS1βPS had elevated levels of basal F-actin. Quantification showed that αPS1 loss caused a similar level of F-actin increase as βPS loss (Fig. 6C,C′,H). Cells lacking αPS1 showed elevated levels of αPS2βPS (Fig. 6D,D′), most likely owing to the lack of competition between the α-subunits to form heterodimers with limited amounts of βPS. Thus, αPS2βPS is able to compensate for the loss of αPS1βPS for stress-fibre organization, but cannot control F-actin levels. We made sure that it was αPS1 loss rather than the resultant increase of αPS2βPS that was elevating F-actin levels, by overexpressing the αPS2 subunit with the Gal4 system. This successfully increased the levels of αPS2βPS, with no change in F-actin levels (Fig. 6E-E″). Absence of αPS1 also resulted in elevated levels of Ena, profilin and Dia, as observed for the absence of βPS (supplementary material Fig. S2) and this was not altered by αPS2 overexpression (data not shown). Conversely, cells lacking profilin still expressed αPS1, showing that the amount of basal F-actin does not influence integrin expression, but that actin fibres are required for normal distribution of focal adhesions (supplementary material Fig. S3). Thus, αPS1βPS is required to control F-actin levels, and αPS2βPS is not able to perform this function. Cells lacking αPS1 do not display cell-shape defects, and therefore F-actin increases are not a secondary consequence of cell-shape changes.

We did not detect any defects in cells lacking αPS2βPS (Fig. 6F,F′). The mutant cells showed elevated levels of αPS1βPS at late stages (Fig. 6G,G′), again suggesting that newly synthesized αPS2 competes with residual expression of αPS1 for limiting amounts of βPS subunit, and therefore that αPS2 expression contributes to αPS1βPS downregulation. Thus, within the limits of our analysis, residual αPS1βPS can substitute for αPS2βPS at late stage 12. Because stress fibres were better organized when αPS1βPS was removed, compared with all integrins removed, this demonstrates that αPS2βPS can organize fibres. We then tested whether the integrin switch per se had an important function for stress-fibre morphogenesis. To do so, we expressed αPS1 in clones of cells lacking αPS2 so that they maintained a high level of αPS1βPS and did not switch to αPS2βPS. These cells had not reoriented the stress fibres at early stage 12 (Fig. 6I-J′″), but recovered and completed reorientation by the end of stage 12 (data not shown). This shows that the switch from αPS1 to αPS2 is essential for the normal rapid return of the stress fibres to their circumferential orientation.

An additional function for αPS2 was suggested by the round egg phenotype caused by clones lacking αPS2 (Bateman et al., 2001) and in the eggs produced by females homozygous for a weak allele of the gene encoding αPS2, if^V2 (supplementary material Fig. S4). However, in our analysis of follicle-cell clones lacking αPS2 we found many examples in which all follicle cells lacked αPS2, yet the oocyte was normally elongated (data not shown). A possible explanation was that αPS2 also functions in the surrounding muscle layer (called the epithelial sheath), which contracts to help oocyte elongation. These muscles are unusual in that they do not fuse, and so clones of mutant cells can be produced (Hudson et al., 2008). We generated clones lacking αPS2 just in these muscle cells and found detachment of the actin from the ends of the muscles (supplementary material Fig. S4), similar to the muscle detachment in embryos (Brabant and Brower, 1993), and this was not rescued by αPS1 (supplementary material Fig. S4), but no round eggs were observed. It might therefore be that the round egg phenotype is caused by loss of αPS2 in both follicle cells and this muscle layer.

In order to define further the contribution of αPS2βPS integrin to stress-fibre organization, we examined a molecular phenotype specific to adhesive structures containing αPS2βPS: the recruitment of tensin.

Follicle-cell adhesions vs focal adhesions in cultured cells: similarities and differences

The changes observed in the follicular epithelium have both similarities and differences with the changes in focal adhesions that occur within vertebrate cells cultured on a 2D ECM. In both cases, distinct adhesive structures arise through a process of maturation. In vertebrate cells, the first structures that form are focal complexes, some of which mature into focal contacts, which in turn can mature into fibrillar adhesions (Zamir et al., 1999). In both vertebrate cells and the follicular epithelium, the adaptor protein tensin is enriched in the most mature structures: fibrillar adhesions and post-stage-11 follicle cells, respectively. Also, in both systems the integrin that forms the connection to the ECM changes. In vertebrate cells this is a change between two fibronectin receptors, αVβ3 and α5β1, whereas in the follicular epithelium the change in integrin implies a change in the ECM ligand from laminin to an RGD-containing ligand (Bunch and Brower, 1992; Gotwals et al., 1994). The main difference is that proteins that are lost in vertebrate cells from the adhesive structures as they mature, such as paxillin and proteins phosphorylated on tyrosine, remain present in late follicle-cell adhesions, coexisting with tensin. Furthermore, both the different integrin structures coexist in vertebrate cells, whereas all adhesions change simultaneously in the follicular epithelium. Finally, the time frame differs, with the transition from a focal complex to a fibrillar adhesion taking approximately 1 hour (Zamir et al., 2000), whereas follicle-cell maturation takes 10.5 hours (Spradling, 1993).

Our findings demonstrate that the composition of adhesive structures does change in normal cells within an intact tissue, and therefore such changes are not just a phenomenon reserved for cells in culture. This indicates that 3D culture does not in all cases represent intact tissues more accurately than 2D culture. The differences between 2D and 3D culture might still highlight an important distinction between integrin adhesion in cells surrounded by ECM compared with those that contact the ECM with part of their cell surface, as epithelial layers do when contacting the basement membrane. It could be that focal-adhesion transitions are restricted to cells in contact with a flat ECM, but to confirm this will require examining integrin adhesions in cells surrounded by an ECM in their native environment.

Integrin switching

Our findings establish a new example of integrin switching during a developmental programme. It is the first example of such a transition in fly tissues, but similar transitions have been observed during the differentiation of mammalian myoblasts and adipocytes, and in C. elegans distal tip cells (reviewed in Meighan and Schwarzbauer, 2008). In all these examples, the developmental switch involves a laminin-binding integrin and an RGD-binding integrin, but the direction of the switch differs. In mammalian cells undergoing differentiation, RGD-binding α5β1 is present in the starting non-differentiated cells and promotes proliferation, and is replaced by laminin-binding α6β1, which promotes differentiation. By contrast, in invertebrates, cells start with laminin-binding αina-1βpat-3/αPS1βPS and switch to αpat-2βpat-3/αPS2βPS. So far, in all cases the switch occurs by reciprocal transcriptional regulation of the two α-subunits. In C. elegans, the single transcription factor vab-3 (Pax6) represses expression of ina-1, while initiating the expression of pat-2 (Meighan and Schwarzbauer, 2007). Early expression of αina-1βpat-3 mediates distal-tip cell migration and its downregulation arrests migration to specify gonad length. Expression of αpat-2βpat-3 turns the gonad, creating organ shape. Our finding that preventing the integrin switch alters the timing of stress-fibre reorientation, rather than stress-fibres structure, suggests that the switch provides robustness to a complicated orchestration of morphogenetic changes, recruitment of intracellular components and development of the egg chamber.

Regulation of the actin cytoskeleton by integrins

Removal of all βPS integrins or just αPS1βPS from follicle cells results in elevation of F-actin and the actin-nucleating proteins Ena, Dia and profilin. With no integrins, the basal actin fibres are completely disorganized, whereas at late stages the actin fibres are well organized in the absence of αPS1βPS. This shows that regulation of actin and actin-nucleating protein levels is independent from organizing stress fibres into parallel arrays.

In the follicular epithelium, cells lacking Ena, profilin or Dia have reduced basal actin (Baum and Perrimon, 2001) (our unpublished results), consistent with a role in stimulating the formation of the stress fibres. In mammalian and follicle cells, Ena localizes at integrin sites and overexpressed Ena causes the formation of actin aggregates (Gertler et al., 1996; Baum and Perrimon, 2001), which we also observed in cells lacking integrins. A link between integrins and the actin machinery was revealed by de novo actin polymerization at integrin adhesions via recruitment of actin-polymerization factors Arp2/3, Ena, profilin and Dia (Chakraborty et al., 1995; Reinhard et al., 1995; Butler et al., 2006). Moroever, profilin co-precipitates with paxillin, indicating that they are in a complex (Mayhew et al., 2006). Consistent with Dia function regulating basal F-actin in follicle cells, stress-fibre formation in cultured cells is mainly mediated by Dia1 and ROCK (Watanabe et al., 1999), and removing ROCK from follicle cells reduces the thickness of stress fibres (our unpublished data). The enhancement of Dia1-induced actin-polymerization by external pulling force (reviewed in Bershadsky et al., 2006) is consistent with the transient increase in Dia protein levels at stage 10B, when stress fibres thicken and the cells flatten, which seems to be regulated by increasing dia mRNA levels from stage 10A (our unpublished results).

Altogether, these data support the view that an increase in actin-nucleating factors could account for the increase in F-actin in follicle cells lacking integrins. Why integrins downregulate the actin machinery while being required for the maintenance of a complex cytoskeletal structure is unknown, but it might be that organizing fibres into parallel arrays works more efficiently if actin polymerization is partially suppressed.

Specific function of αPS2 and tensin recruitment

The specificity of function between αPS1 and αPS2 was previously shown to be dependent on their extracellular domains (Martin-Bermudo et al., 1997). We had interpreted this to show that the important property that individual α-subunits bring to the heterodimer is specific interaction with extracellular ligands. In addition, we have now found that extracellular specificity also changes intracellular events. This raises the key issue of how the extracellular (and/or transmembrane) domain could specify the recruitment of an intracellular protein.

We can envision three possible explanations: (1) the different ECM interaction alters the ability of the integrin to resist force. Actomyosin-driven unfolding of proteins bound to the integrin might expose new interaction sites (e.g. del Rio et al., 2009). The specific integrin-ECM interaction could differ in ligand-binding strength or matrix rigidity, affecting the ability to resist force, and thus the level of new site exposure. Supporting this, increasing ECM rigidity strengthened the integrin-cytoskeleton link in fibroblasts (Choquet et al., 1997). Thus, the interaction of αPS2βPS with its ligand might be able to resist more force than αPS1βPS, thus exposing tensin recruitment sites. The proposed role of tensin in α5β1 translocation from focal contacts to fibrillar adhesions and fibronectin fibril formation (Pankov et al., 2000) suggests that tensin recruitment feeds back to increase matrix rigidity. (2) The different ECM interactions affect the density of integrin clusters, which affects tensin recruitment. The density of αPS2βPS clusters might be higher than αPS1βPS clusters, providing higher avidity for a low-affinity interaction with tensin. (3) Recruitment or activation of another transmembrane protein in the same membrane, which in turn recruits tensin through its intracellular domain. For example, specific integrin heterodimers bind tetraspanins, which in turn bind protein kinase C, bringing it into proximity with integrins (Zhang et al., 2001). In another example, a combination of signals from α5β1 and the proteoglycan syndecan-4, which both bind extracellular fibronectin, leads to the activation of p190RhoGAP and the suppression of Rho (Bass et al., 2008). Thus, switching the integrin extracellular binding might bring the integrin adjacent to other receptors that synergize to recruit tensin.

Immunostainings

Antibody stainings were carried out on female ovaries dissected in PBS, fixed 10 minutes in 4% formaldehyde, permeabilized, blocked for 1 hour in PBS 0.3% Triton X-100, 0.5% BSA, and stained overnight at 4°C for both primary and secondary antibodies. The following antibodies were used (HB stands for The Developmental Studies Hybridoma Bank): anti-βPS [HB CF.6G11 mouse monoclonal antibody (MmAb); 1:100] (Brower et al., 1984), anti-αPS1 (HB DK.1A4 MmAb; 1:10 (Brower et al., 1984), anti-αPS2 (5D6 rat mAb; 1:10) (Bogaert et al., 1987), anti-αPS3 [rabbit polyclonal antibody (RpAb); 1:100] (Grotewiel et al., 1998), anti-αPS4 (RpAb; 1:200) (Krzemien et al., 2007), anti-talin (E16B; MmAb; 1:50) (Brown et al., 2002), anti-PINCH (RpAb; 1:500) (Clark et al., 2003), anti-_PY (4G10 MmAb; 1:500; Upstate #05-321), anti-pFAK^Y397 (RpAb; 1:500; Biosource), anti-FAK (RpAb; 1:500) (Palmer et al., 1999), anti-PAK (RpAb; 1:1000) (Harden et al., 1996), anti-Zipper (RpAb; 1:500) (Jordan and Karess, 1997), anti-profilin (HB ch1J MmAb; 1:50) (Verheyen and Cooley, 1994), anti-Ena (HB 5G2 MmAb; 1:10) (Bashaw et al., 2000), anti-Dia (RpAb; 1:100) (Afshar et al., 2000), anti-GFP (RpAb; 1:500; Abcam Ab290), anti-GFP (MmAb; 1:500; Roche), anti-paxillin (RpAb; 1:250) (Chen et al., 2005), anti-Wech (RpAb; 1:20) (Loer et al., 2008), anti-Eyeless (RpAb; 1:200) (Adachi et al., 2003), anti-Twin-of-eyeless (RpAb; 1:500) (Jacobsson et al., 2009). To visualize tensin and ILK we used GFP-tagged genomic rescue constructs (Zervas et al., 2001; Torgler et al., 2004) and Zasp, a GFP gene trap (G00189) (Jani and Schock, 2007).

Secondary antibodies were Alexa-Fluor-488-conjugated anti-rabbit and anti-mouse from Molecular Probes, and Cy5-conjugated anti-rabbit, anti-mouse and anti-rat from Jackson Laboratories, all used at 1:200. F-actin was stained using rhodamine-phalloidin (#R-415, Molecular Probes, 1:500). The DNAse-I staining was performed using a DNAseI–Alexa-Fluor-488 conjugate (0.3 μM, Molecular Probes). Samples were mounted in Vectashield H-1000 or Vectashield H-1200 with DAPI (Vector Laboratories). Confocal images from an Olympus Fluoview FV1000 were assembled with Adobe Photoshop CS3.

In situ hybridization on ovaries

In situ hybridization was performed as in Palacios and St Johnston (Palacios and St Johnston, 2002) with antisense DIG probes against fragments of mew and if mRNA, and pax6 orthologue mRNAs were transcribed from DGRC cDNAs: GH01157 (ey), GH14454 (toy), AT09010 (eg), GH22493 (toe). Ovaries were mounted in 1:1 PBS/glycerol and imaged using a LEICA DMR microscope and an Optronics digital camera.

FISH was performed to detect profilin and dia mRNA using antisense DIG probes transcribed from DGRC cDNAs LD15581 and LD14246, respectively. Hybridized ovaries were incubated for 2 hours with an anti-GFP to detect mys^XG43 clones and an anti-DIG-HRP (Roche, 1:2000) followed by amplification with Cy5 tyramide reagent (Perkin Elmer).