Selection of mutations to disrupt the talin ABS domains

Several biochemical studies have defined and characterized the ABS2 and ABS3 domains of Talin [17,22–24,31]. The ABS2 domain extends over a sizable region of the Talin Rod, over 600 amino acids long, as it involves two different helical bundles, R4 and R8, that are separated by 3 domains and ~650 amino acids (Fig 1A) [6,31]. The R4 and R8 region share some similarity with R13 which encompasses ABS3 and contain a series of highly conserved surface residues that were identified as important for binding to actin. Mutating three of these conserved surface residues in R4 (K922, Q923, Q930 in vertebrate talin), and three additional residues in R8 (K1500, R1510, and K1522 in vertebrate talin) to glutamic acid residues reduced the actin binding ability of talin, as judged by in vitro actin sedimentation assays, by over 60% [31]. These sites are conserved in Drosophila and correspond to K941, Q942, Q949 in R4 and K1510, R1520, K1532 in R8 of Drosophila talin, respectively (Fig 1B and 1C). To disrupt ABS2 we mutated these 6 residues to glutamic acid in Drosophila talin and confirmed using a series of biochemical experiments that introducing these mutations did not impact protein folding or stability. To disrupt ABS3 the conserved residues K2450, V2451, K2452 were mutated to aspartic acid, a change that has been shown by us and others to greatly reduce, though probably not eliminate, actin binding through ABS3 but not impact protein folding or stability [17,23,24,31]. Previous analysis of the KVK/DDD ABS3 mutants [17,24] found anywhere between 40%-73% decrease in actin binding in an in vitro actin co-sedimentation assay. These in vitro experiments likely represent an overestimate of the actin binding activity of these mutations as they are designed to optimize actin binding. To test the structural integrity of the mutated domains using Nuclear Magnetic Resonance (NMR) and Circular Dichroism (CD), the wildtype R4, R8 and R13 domains, and the corresponding mutated R4 domain (R4_3E; K941, Q942, Q949), R7/R8 domains (R7R8_3E; K1510, R1520, K1532) and R13 domain (R13_3E; K2650, V2651, K2652) were each recombinantly expressed and purified (see materials and methods). The 1H,15N HSQC spectra of all three wildtype domains show good dispersion and uniform, peak distribution confirming that the fly rod domains are correctly folded (S1A Fig). Similarly, the CD spectra confirmed that all three rod domains were helical in nature (S1D–S1F Fig). The mutant proteins all gave HSQC and CD spectra that were similar to the corresponding wildtype versions confirming that the mutations to the exposed surface residues did not perturb domain folding or stability, similar to what was shown previously for equivalent mutations in mouse talin [22, 31]. Having validated the mutants did not affect the structural integrity of the fly Talin domains, we next generated fly mutant lines containing the 6 point mutations in ABS2 (K941E, Q942E, Q949E, K1510E, R1520E, K1532E), referred to henceforth as rhea^ABS2 allele or ABS2 mutant and the 3 point mutations in ABS3 (K2450D, V2451D, K2452D), referred to henceforth as rhea^ABS3 allele or ABS3 mutant (see materials and methods, Fig 1A–1C).

Generation of ABS mutations using CRISPR

To introduce the ABS2 and ABS3 mutations into rhea, the gene that encodes talin in flies, we used a CRISPR/Cas9-based approach. Two different CRISPR/Cas9-mediated methodologies were used to genetically engineer the endogenous rhea locus, a single-stranded oligodeoxynucleotides (ssODNs) approach was used to introduce mutations into the ABS3 domain while a strategy of “scarless” homology-directed repair (HDR) with a double-stranded DNA (dsDNA) donor template (see materials and methods, [47]) was used to alter the ABS2 domain (Fig 1D and 1E). Briefly, to modify the ABS2 sequence two guide RNAs (gRNAs) that targeted sites 5’ and 3’ of the sequence corresponding to the ABS2 domain were used as well as a dsDNA donor construct. The donor construct featured homology arms containing approximately 2kb of sequence flanking a visible selection marker, DsRed, that contained PBac transposase target sites allowing excision (see materials and methods, Fig 1D). Each of the two homology arms contained a cluster of mutations in the ABS2 site, namely K940D/Q941D/Q948D in R4 and K1510E/R1520E/K1532E in R8. In the first step of mutagenesis the DsRed marker was used to identify successful editing events while in the second step this DsRed marker was removed by crossing to a PBac transposase source. To modify the ABS3, a ssODN donor of 200 base pairs long was synthesized that contained the K2670D/V2671D/K2672D mutations (Fig 1E). This was followed by a genetic and PCR based screening approach to identify candidate mutant lines (see materials and methods). Following the identification of multiple candidates, ABS2 and ABS3 mutant lines were sequenced extensively both upstream and downstream of the mutation sites to confirm the presence of the mutation and to ensure no other deleterious events took place at the rhea locus (Fig 1F and 1G). Using the approach detailed here we successfully generated multiple independent mutated lines of ABS2 and ABS3. As the different ABS2 and ABS3 mutant lines displayed identical sequencing results, phenotypic analysis was then performed on only 1 line per ABS mutation to investigate their role.

Actin binding through the talin ABSs is required for viability

We noted that the ABS2 (rhea^ABS2/ABS2) or ABS3 (rhea^ABS3/ABS3) mutant flies were non-viable and failed to complement null alleles of rhea. Moreover, introducing a ubiquitously expressing talin rescue construct (ubi::TalinGFP, [48]) into the background of null alleles of rhea, or homozygous mutants of the ABS2 or ABS3 mutant alleles rescued the embryonic lethality associated with loss of talin to the same extent, specifically, until pupation (Fig 2A). These data are consistent with ABS2 or ABS3 mutant alleles acting like genetic loss-of-function alleles of rhea. Further analysis of the stage at which lethality took place in the different mutants showed that while homozygous null mutants of rhea or homozygous ABS2 mutant flies died during the first instar larval stage, homozygous ABS3 mutants died in the second instar larval stage and displayed significantly higher L1 survival rate than ABS2 mutants (Fig 2A and 2B). This suggested that the phenotype of the ABS2 mutant allele was similar to that of a null allele of rhea while the phenotype of the ABS3 mutant allele was not as strong. Importantly, when we generated transheterozygotes, flies that contain both the ABS2 and ABS3 mutant alleles but do not produce wildtype talin (rhea^ABS2/ABS3) (see materials and methods), the flies exhibited a much weaker phenotype than either ABS mutants by themselves (Fig 2A and 2B). Specifically, ABS2/ABS3 transheterozygote mutant flies survived until pupal stages similarly to talin mutants expressing the talin rescue construct. To learn more, we employed a more sensitive method to compare the strength of the phenotypes of talin mutants. In particular, we measured “motility” by quantifying the percentage of fly larvae that were able to move more than 5 mm over the course of at least 1 hour, (Fig 2C–2H see materials and methods). This analysis showed that null mutant flies which lack all talin function exhibit almost no movement, with only about 4% of larva moving at least 5 mm during the given time period compared to 100% of wildtype flies. Flies homozygous for the ABS2 mutant allele exhibit a somewhat weaker phenotype as 25% moved at least 5 mm, while for the ABS3 mutant that proportion increased to 72%. In comparison nearly 88% of ABS2/ABS3 transheterozygotes flies exhibited such motility. To determine whether these mobility phenotypes corresponded to visible defects in muscles we introduced actinin::GFP, a marker for sarcomeric architecture and muscle attachment, into the background of our mutants (Fig 2I–2L). Mutants were analyzed in the latest larval stage that they were able to reach, and although null alleles of talin and ABS2 mutants proved too fragile for the staining protocol, we were able to image second instar larva from ABS3 mutants and third instar larva from ABS2/ABS3 transheterozygotes mutants. Surprisingly, we observed no noticeable muscle defects in ABS3 mutants or ABS2/ABS3 transheterozygotes suggesting alternative causes for their motility phenotype, such as neuronal dysfunction. These data are consistent with the interpretation that the ABS2 mutant allele is a strong loss of function allele that is only slightly less strong than a complete null, the phenotype of the ABS3 mutant allele is intermediate, while ABS2/ABS3 transheterozygotes exhibit only a mild phenotype.

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Fig 2. Viability and motility of talin ABS2 and ABS3 mutants.

(A) Viability of ABS2 and ABS3 homozygote mutants (rhea^ABS2/ABS2 and rhea^ABS3/ABS3 respectively) compared to wildtype (Wt) (rhea^+/+) and null (rhea^-/-) flies. Talin null allele does not complement either ABS mutants but transheterozygote (rhea^ABS2/ABS3) flies display longer viability similar to mutant flies rescued by the expression of a Wt talin (+TalinGFP). (B) L1 Survival rate. Percentage of L1 larvae successfully hatching 12–24 hours after egg deposition (AED). (C) L1 motility. Percentage of L1 larvae capable of moving at least 5 mm away from hatching site. (D-H) Representative picture of L1 larvae from each genotype. (I-L) Confocal images of whole mount L2 (I, J) and L3 (K, L) larvae of the indicated genotype expressing α-actinin-GFP to visualize muscle structure. Survival and motility rates compared using chi-square test: * P<0.05; NS, no significance; all genotypes n>100. Scale bars: 1 mm.

https://doi.org/10.1371/journal.pgen.1011224.g002

The talin ABSs are required for multiple aspects of embryonic development

Complete loss of talin in embryos, achieved by using genetic tools to remove both the maternal as well as zygotic expression of talin (see materials and methods), results in strong embryonic defects that impact multiple tissues and processes [15]. Notable among these are disruption of long-lasting stable attachments of muscle cells to tendon cells, and the failure of two dynamic morphogenetic processes, germband retraction (GBR) and dorsal closure (DC). The attachment of muscles to tendons occurs via integrin-based adhesions at myotendinous junctions (MTJs) that form in late fly embryogenesis [49]. Germband retraction involves the pulling back of the caudal end of the embryo, which at stage 12 is looped back so that it is near the head, such that it reaches its final position at the posterior of the embryo [50]. Dorsal Closure is a morphogenetic process in which a large hole on the dorsal side of the embryo is enclosed by the movement of two lateral sheets of epithelia [36,50]. Fly embryos that completely lack talin (rhea^-/-) are characterized by a high proportion of embryos that exhibit an abnormal folded appearance due to failed GBR, a high proportion of embryos that exhibit a large dorsal hole due to failed DC, and a large proportion of embryonic muscles that are detached and rounded up due to failed attachment of muscles to MTJs (Fig 3A and 3B; quantified in I-K). Flies that completely lack wildtype talin but contain the talin ABS2 mutant allele exhibited a high penetrance of DC and GBR defects, similar to the talin null allele, but showed a lower proportion of detached muscles compared to the null allele (Fig 3C; quantified in 3I–3K). Flies that completely lack wildtype talin but contain the ABS3 mutant allele also exhibited a high penetrance of DC and GBR defects, similar to the null allele, but a much lower proportion of detached muscles compared to the null allele (Fig 3D; quantified in 3I–3K). In contrast flies that completely lack wildtype talin but contain both the ABS2 and ABS3 mutant alleles exhibited a much lower penetrance of DC and GBR defects relative to the null allele, as well as a much lower proportion of detached muscles (Fig 3E; quantified in 3I–3K). Assaying a non-embryonic integrin-associated phenotype, the formation of blisters in the fly wings due to loss of apposition of the two layers of the fly wing, revealed similarly strong phenotype in rhea null, ABS2, and ABS3 mosaic mutants (Fig 3H, see materials and methods). Taken together these data show that the talin ABS sites play diverse, essential roles in fly development and tissue maintenance, and, as revealed by the transheterozygotes, they share a great deal of functional redundancy. Nevertheless, in some processes, such as in muscle attachment, disruption of ABS2 causes stronger phenotype than disruption of ABS3, suggesting it has a broader role.

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Fig 3. Talin ABS requirement for muscle and embryonic development.

(A-G) Confocal images of whole-mount embryos stained with muscle cytoskeleton marker Myosin Heavy Chain (MHC) at stage 17 (scale bar: 50 μm). (H-J) Penetrance of wing blisters (H), dorsal closure (I) and germband retraction (J) defects in embryos of the indicated genotypes. Dorsal closure and germband retraction defects of rhea^ABS2/ABS3 compared to rhea^ABS3/ABS3 using chi-square test: *P<0.05; all genotypes n>30. (K) Number of detached muscles per embryo. Floating bar chart representing minimum and maximum value per genotype (n>10). Mean value represented by a solid line. Each lower-case letter (a-g) indicates a different statistical group where two different letters represent statistically different results (P<0.05) using one-way ANOVA, Dunnett’s post-hoc, P<0.05.

https://doi.org/10.1371/journal.pgen.1011224.g003

ABS2 but not ABS3 is required for talin recruitment to sites of adhesion

The null-like embryonic phenotypes observed in flies expressing only the ABS2 mutant allele were consistent with a strong disruption to integrin-mediated cell-ECM adhesion. Such a defect could be caused by failure to express talin. To test if there were differences in the expression of the ABS mutations we carried out qPCR experiments which showed that both the ABS2 and ABS3 mutant alleles were expressed at similar levels to that of wildtype talin (Fig 4L). Since qPCR measures transcription but not translation or protein stability we used immunohistochemistry to analyze Talin protein levels at sites of adhesion in MTJs (quantified in Fig 4K). To assess the ability of the ABS2 mutant alleles to support recruitment and/or maintenance of talin to MTJs compared to the null and ABS3 mutant alleles of talin we took advantage of a previously described methodology (49). In this approach integrin, which localises to muscle ends independently of talin, is used to label MTJs allowing the determination of the ratio of talin found at MTJ compared to talin found in the cytoplasm (Fig 4A) (49). In a wildtype embryonic muscle, talin is enriched at MTJs by approximately five-fold compared to the cytoplasm (Fig 4K). In embryos lacking any talin expression (rhea^-/-), muscles detached from the ECM but MTJs were still identifiable by integrin labeling allowing us to observe that talin was not enriched at MTJs (Fig 4B, the 1:1 ratio of intensity seen in Fig 4K is consistent with a lack of talin enrichment). In heterozygous embryos that contained only one copy of the wildtype talin allele (rhea^-/+) levels of talin were reduced by about half in MTJs compared to fully wildtype controls (rhea^+/+) (Fig 4C and 4K). In embryos that expressed only the ABS2 mutant allele (rhea^ABS2/ABS2) muscle detachment was observed and, as in the null allele, integrin but not talin was recruited to MTJs (Fig 4D, and 4K). Finally, in embryos that expressed only the ABS3 mutant allele (rhea^ABS3/ABS3), talin was recruited to MTJs to a similar degree as in wildtype controls (Fig 4F and 4K). Overall, these results suggest that the ABS3 domain of talin is dispensable, while the ABS2 domain of talin is at least partially required, for talin recruitment to, or maintenance at, sites of adhesion. These results however left open the possibility that the ABS2 allele interfered with talin expression by impinging on the translation or stability of the talin protein.

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Fig 4. ABS2, but not ABS3 is required for talin recruitment to sites of adhesion.

(A-J′′) Talin recruitment at MTJs. Confocal z-stacks of MTJs and surrounding muscles in stage 17 embryos stained for MHC (white), the integrin subunit αPS2 (magenta) and Talin (green). Scale bar: 25 μm. MTJs location was determined by αPS2 staining in between muscle cells (arrowheads in A-I). (K) Relative localization of Talin to MTJs. Mean value represented by solid line, error bar show s.e.m. and dashed line indicates the absence of recruitment (n>15 per genotype). Each lower-case letter (a-c) indicates a different statistical group (P<0.05) using one-way ANOVA, Dunnett’s post-hoc. (L) qPCR analysis of rhea^+/−, rhea^+/ABS2 and rhea^+/ABS3 heterozygote flies compared to wildtype (Wt) flies (two-tailed Student’s t-test, *P<0.05). Error bars represent s.e.m.

https://doi.org/10.1371/journal.pgen.1011224.g004

Further insight into the question of whether the ABS2 allele interfered with protein expression emerged in experiments that measured talin levels at MTJ when the mutant ABS3 allele was combined either with a null or an ABS2 allele. In a mutant talin ABS3 heterozygotes, meaning a background that contains a talin null allele and an ABS3 mutant allele (rhea^ABS3/-) the recruitment of the ABS3 mutant talin protein to sites of adhesion at MTJs was reduced by half compared to a wildtype (rhea^+/+) (Fig 4G and 4K). This is similar to what is observed in heterozygous null embryos that have one copy of wildtype talin and one copy of a null allele of talin (rhea^+/-; Fig 4C and 4K). Such results were consistent with the idea that the mutant talin ABS3 protein was recruited with similar efficiency as the wildtype talin protein to MTJs. Importantly, in a background that contained one copy of the ABS3 and one copy of the ABS2 mutant alleles (rhea^ABS2/ABS3) the levels of talin recruited to MTJs were similar to those seen in wildtype controls (Fig 4H and 4K). These results are consistent with the interpretation that the talin protein with the ABS2 mutation is abundantly expressed and can be visualized at the MTJs if it receives assistance in reaching the membrane, as it does in the presence of ABS3.

We sought to test the idea that a main causative defect in ABS2 mutants was their failure to localize to, or be retained at, the membrane, and that this defect could be overcome by synergistic interactions with ABS3 mutants [51]. To this end we used an allele of talin, rhea^K17E, that contains a point mutation that interferes with the direct binding of talin to Rap1, an interaction that we showed previously does not impact the expression of talin but disrupts the proper recruitment and/or retention of talin to the membrane [43]. Consequently, homozygous rhea^K17E mutants behave like talin null mutants, even though the talin protein is made in wildtype amounts. Our hypothesis predicted that, unlike ABS3 mutants, rhea^K17E and rhea^ABS2 mutants share a molecular defect in terms of recruitment. For example, as we previously showed for rhea^K17E heterozygotes (rhea^K17E/+) [43], in an embryo containing one copy of wildtype talin and one copy of talin with the ABS2 mutations (rhea^ABS2/+) we observed levels of talin at sites of integrin-mediated adhesion at MTJs that were slightly lower than, but near to, those observed in a wildtype embryo (rhea^+/+; Fig 4A, 4I and 4K). In comparison, in an embryo containing one copy of the rhea^K17E allele and one copy of talin with the ABS2 mutations (rhea^ABS2/K17E) we observed loss of localization to MTJs to a similar extent as we saw in talin null embryos (rhea^-/-; Fig 4B, 4J and 4K). Furthermore, the rhea^K17E allele, much like the talin null allele, fails to functionally complement the ABS2 mutation. In particular, embryos lacking wildtype talin but containing one copy of the rhea^K17E allele and one copy of the ABS2 mutant talin allele (rhea^ABS2/K17E) showed phenotypes that were similar to those observed when the ABS2 mutant allele was combined with a talin null allele (rhea^ABS2/-) (Fig 3F and 3K and 3G and 3K for the ABS2 and ABS3 mutant alleles, respectively). The failure of the rhea^K17E allele to complement or rescue flies containing the ABS2 mutation supports the interpretation that these alleles share a functional defect. In contrast, in the presence of a wildtype allele or an ABS3 mutant allele (rhea^ABS2/+ or rhea^ABS2/ABS3), a talin protein containing the ABS2 mutation can be recruited stably to sites of adhesion. Based on these data we propose that the ABS2 domain, but not the ABS3 domain, of talin plays an important role in talin recruitment to the membrane.

Actin organization at MTJs is disrupted in talin ABS2 mutants but not talin ABS3 mutants

In late-stage fly embryos, actin is enriched at the MTJs (Fig 5A; [17]). Actin enrichment can be quantified by staining actin in muscles using phalloidin and determining the ratio of the intensity of actin staining in the cytoplasm to the actin staining intensity at the MTJs (Fig 5F). In an embryo that lacks talin, actin was not enriched at muscle ends, even in MTJs that remain attached (Fig 5B and 5F; [17]). A similar, but less severe, phenotype of loss of actin enrichment was observed in embryos that expressed only the ABS2 mutant of talin (rhea^ABS2/ABS2; Fig 5C and 5F). In comparison, in embryos that expressed only the ABS3 mutant (rhea^ABS3/ABS3) the levels of actin at muscle ends were similar to those seen in wildtype controls (Fig 5D and 5F). Similarly, in embryos that lacked wildtype talin but contained one copy of the ABS2 allele and one copy of the ABS3 mutant allele (rhea^ABS2/ABS3) the levels of actin at muscle ends were also comparable to wildtype controls (Fig 5E and 5F). Although the poor recruitment and/or maintenance of the talin protein with the ABS2 mutation complicates the interpretation of the actin phenotype of this allele, these data suggest that a talin containing the ABS3 mutation retains some ability to recruit and organize actin at sites of adhesion at muscle ends. Taken together with the near wildtype rescue of actin organization in the ABS2 and ABS3 transheterozygote mutants this data is consistent with a redundancy between the two ABSs in actin organization.

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Fig 5. Actin organization at MTJs is disrupted in ABS2 mutants but not in ABS3 mutants.

(A-E′) Confocal z-stacks of stage 17 embryos co-stained with Rhodamine-Phalloidin to label F-actin (white) and the integrin subunit αPS2 (magenta). MTJs location determined by integrin staining (arrowheads in A-E′). Scale bar: 25 μm. (F) Relative localization of F-actin to MTJs. Mean value represented by solid line, error bars show s.e.m. and dashed line indicates the absence of recruitment (n>30 per genotype). Lower-case letter (a-c) indicates a different statistical group (P<0.05) using one-way ANOVA, Dunnett’s post-hoc.

https://doi.org/10.1371/journal.pgen.1011224.g005

ABS2 and ABS3 have distinct as well as overlapping roles in recruiting the integrin adhesion complex

Integrin-mediated adhesion is dependent on both the ability of integrins to establish and maintain a multiprotein complex inside the cell, and to attach to ECM molecules outside of the cell [52,53]. Talin is essential for both of these processes as it is required for the assembly of the intracellular integrin adhesion complex and also for the attachment of integrins to their ECM ligands [15,32,54,55]. To determine whether the defects observed in the ABS mutants result from disruption of either, or both, of these processes, markers for the integrin adhesion complex and the ECM were analyzed. To visualize the intracellular integrin adhesion complex in wildtype and mutant embryos two different markers were used, paxillin [56] and PINCH [57] (Fig 6A–6L). The relative localization of paxillin and PINCH to sites of adhesion at the MTJs was quantified by determining the ratio of the intensity of paxillin or PINCH staining in the cytoplasm to their staining intensity at MTJs (Fig 6K and 6L, respectively; see materials and methods). In wildtype muscles, both paxillin and PINCH were significantly enriched at the muscle ends compared to the cytoplasm (Fig 6A and 6B; quantified in K&L respectively). This enrichment to muscle ends was not observed in embryos lacking talin, as both Paxillin and PINCH failed to concentrate at the MTJs resulting in a 1:1 ratio of cytoplasmic to MTJ staining intensity (Fig 6C and 6D; quantified in K&L respectively). A similar failure to concentrate at the MTJs was observed for either paxillin or PINCH in embryos that lacked endogenous talin but expressed the ABS2 mutant (rhea^ABS2/ABS2; Fig 6E and 6F; quantified in 6K and 6L, respectively). However, near wildtype levels of enrichment at the MTJs were seen in embryos that lacked endogenous talin but expressed the ABS3 mutant (rhea^ABS3/ABS3; Fig 6G and 6H; quantified in 6K and 6L respectively). Notably, in embryos that lacked wildtype talin but contained one copy of the ABS2 and ABS3 mutant alleles (rhea^ABS2/ABS3) paxillin levels, but not PINCH levels, at the MTJs were partially reduced compared to wildtype control (rhea^ABS2/ABS3; Fig 6I and 6J; quantified in 6K and 6L respectively). These results suggest that Paxillin recruitment to sites of integrin-mediated adhesion is dependent on ABS2 function to a greater extent than PINCH recruitment, while ABS3 is not required for the recruitment of either paxillin or PINCH.

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Fig 6. ABS2 and ABS3 roles in recruiting the integrin adhesion complex and attachment to the ECM ligand Tiggrin.

(A–J) Confocal z-stacks of MTJs in stage 17 embryos stained for IAC components paxillin and PINCH (green, A-J), the integrin subunit αPS2 (magenta, A-J and green, M—O′′). (K&L) Relative localization of Paxillin (K) and PINCH (L) to MTJs localized by αPS2 staining. Mean value represented by solid line, error bars show s.e.m. and dashed line indicates the absence of recruitment (n>15 per genotype). Lower-case letter (a-d) indicates a different statistical group (P<0.05) using one-way ANOVA, Dunnett’s post-hoc. (M-O′′) Confocal z-stacks of MTJs in stage 17 embryos stained for the ECM ligand tiggrin (magenta, M′-O′′). (P) Pixel intensity profiles of tiggrin (top panel Wt, middle panel rhea^ABS2/ABS2 and lower panel rhea^ABS3/ABS3) across the widths of the boxed areas indicated in the corresponding images. Pearson correlation coefficient (r) > 0.7 for all genotypes. Scale bar: 25 μm.

https://doi.org/10.1371/journal.pgen.1011224.g006

To test whether the talin ABSs were required for integrin-mediated attachment to the ECM we visualized the staining of the ECM ligand Tiggrin at muscle ends. In wildtype MTJs, Tiggrin and αPS2-integrin localization at MTJs overlapped (Fig 6M; visualized using a staining intensity profile in P). It has been previously shown that mutations that disrupt integrin-mediated attachment to the ECM resulted in clear separation of Tiggrin from the integrin at MTJs [42, 48]. However, in embryos that lacked endogenous talin but expressed either the ABS2 or ABS3 mutant talin, Tiggrin and αPS2-integrin localization at MTJs still overlapped (Fig 6N and 6O, respectively; visualized using a staining intensity profile in P). These results suggest that neither ABS2 or ABS3 are required for establishing or maintaining the linkage of integrins to their ECM ligands.

Vinculin overexpression ameliorates the phenotype of talin ABS3 domain mutants

One of the main proposed functions of the talin ABSs is to facilitate force transmission across the protein by linking to actin. Force transmission across the talin protein is thought to lead to stereotypic unfolding of the repeating helical bundles that constitute the large rod domain of talin, a conformational change that exposes cryptic protein-protein binding sites [23,27–29,31,58]. In particular, talin contains a large number of binding sites in its rod domain for the scaffolding protein vinculin [19,23,59,60] that are not available in the folded conformation but are available in the unfolded conformation [27,29,61,62]. It has been proposed that the exposure of these cryptic binding sites in talin and the subsequent binding of vinculin to talin plays an important role in the consolidation and maturation of integrin-based adhesions [30,31,62]. According to this proposed model, the binding of actin through ABS3 allows actin to pull on talin and expose cryptic vinculin binding sites that, when bound by vinculin, promote actin binding by ABS2 and adhesion stabilization. Key evidence in support of this model was provided by experiments showing that expression of an activated vinculin can compensate for loss of actin binding through ABS3 [23]. To reproduce this experiment in flies we drove ectopic vinculin expression in our ABS2 and ABS3 mutant alleles. We are not able to use activated vinculin since it has been shown that, in flies, its expression has dominant negative effects resulting in loss of function phenotypes in integrin-mediated adhesion [63]. Instead, we employed overexpression of wildype vinculin in muscles of zygotic talin mutants, that contain maternal talin and therefore have weaker phenotypes than the complete loss of talin, using vinculin overexpression in wildtype flies as the control. Previous work characterizing vinculin overexpression suggested that expression of constitutively active vinculin, but not overexpression of wildtype vinculin, had a severe impact on viability (63). In contrast, we noted that overexpression of wildtype vinculin also resulted in a decrease in viability of about 30% across all genotypes (Fig 7B). Nonetheless, we were able to assess the impact of overexpression of vinculin in the background of the different talin mutants. In combination with the talin null allele, or an ABS2 mutant allele (rhea^-/- or rhea^ABS2/ABS2) we noted had no impact on the phenotype (Fig 7A–7C). In comparison, overexpression of vinculin in the background of a talin ABS3 mutant allele (rhea^ABS3/ABS3) substantially ameliorated the phenotype as some flies were able to survive longer, and equalized larval mobility between ABS3 mutants and control wildtype larvae (Fig 7A–7C). These results suggest that overexpression of vinculin, despite having a slight negative impact on viability, can nevertheless ameliorate the phenotype caused by disruption of the ABS3 domain of talin.

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Fig 7. Vinculin overexpression partially rescues talin ABS3 mutants’ viability phenotype.

(A) Viability of Mef2::GAL4 flies with or without a UAS promoter expressing vinculin-TagRFP in genetic backgrounds as shown. (B) L1 Survival rate. Percentage of L1 larvae successfully hatching 12–24 hours after egg deposition (AED). (C) L1 motility. Percentage of L1 larvae capable of moving at least 5 mm away from hatching site. (D-H) Muscles in live wildtype stage 17 embryos expressing the TagRFP -tagged vinculin (scale bar: 25 μm). (I) Relative localization of vinculin- TagRFP to MTJs in live wildtype stage 17 embryos expressing the TagRFP -tagged vinculin. (J-N) Native vinculin expression in stage 17 embryos using genomically GFP-tagged vinculin (Vinculin::GFP). (O) Relative localization of Vinculin;;GFP to MTJs in live wildtype stage 17 embryos expressing. Mean value represented by solid line, error bars show s.e.m. and dashed line indicates the absence of recruitment (n>15 per genotype). Lower-case letter (a-c) indicates a different statistical group (P<0.05) using one-way ANOVA, Dunnett’s post-hoc.

https://doi.org/10.1371/journal.pgen.1011224.g007

We also noted in these experiments that the phenotype observed in the ABS3 mutant background where vinculin is overexpressed was, in terms of mobility, and viability, indistinguishable from what was observed in ABS2/ABS3 transheterozygotes mutants (Fig 7A–7C). In addition, overexpression of vinculin did not modify the phenotype of ABS2/ABS3 transheterozygotes mutants (Fig 7A–7C). These data implied that the synergistic interaction between the ABS2 and ABS3 mutant talin proteins could be mimicked by overexpressing vinculin. Further support for this idea came from experiments studying vinculin recruitment in talin mutants. Vinculin recruitment was analyzed using two different approaches: by ectopic expression of a RFP tagged version of Vinculin that was used in the overexpression experiments (Vinculin-TagRFP; (63)) or by analyzing the distribution of a genomically GFP-tagged (Vinculin::GFP; [64]), allele of vinculin Both approaches showed that in a talin null background vinculin recruitment to sites of adhesion at muscle attachment sites was severely curtailed compared to wildtype controls (Fig 7D, 7E and 7J, 7K, quantified in 7I). This defect in vinculin recruitment was nearly as severe in talin ABS2 mutants (Fig 7F and 7L, quantified in 7I). However, in talin ABS3 mutants or in transheterozygote mutants vinculin recruitment was substantially better compared to the null or ABS2 mutants (Fig 7G, 7H and 7M, 7N, quantified in 7I). The observation that ABS3 mutants retained some capacity to support vinculin recruitment to sites of adhesion helps explain why they are amenable to rescue by vinculin overexpression. This result suggests that the general relationship between talin ABS3 mutants and vinculin activity is conserved between mammals and flies.

In vivo evidence for the functional importance of talin dimerization

Talin has long been thought to form a dimer [24,65–68] but this has not been demonstrated in a functional capacity in vivo before. The substantial rescue, in terms of both talin recruitment and developmental phenotypes, that was observed by combining the talin ABS2 and ABS3 mutants suggested the existence of a talin dimer that contained one ABS2 and one ABS3 protein. Specifically, we hypothesized that a talin dimer composed of one ABS2 and one ABS3 mutant talin proteins could perform more functions than a talin dimer composed of two ABS2 or two ABS3 mutant talin proteins. For example, we hypothesized that while a talin dimer composed of two mutant ABS2 proteins was not recruited to sites of adhesion and was unable to function in adhesion, a dimer composed of one ABS2 and one ABS3 mutant was recruited to, and was partially functional at, sites of adhesion (Fig 4K). To directly test this hypothesis, we utilized the previously described R2531G talin mutant, that is known to block talin dimerization [17]. In addition to blocking dimerization the R2531G mutation disrupts the function of the ABS3 domain [17,24]. This allowed us to compare the impact of a simple disruption in ABS3 versus disruption of ABS3 coupled with an inability to form dimers. For this experiment we relied on two transgenes we previously generated and characterized that expressed, under the control of ubiquitous promoter, a talin rescue transgene containing either the ABS3 or the R2531G mutations ([17]; Fig 8A–8C; Quantified in J). As shown in our previous study, while introducing a wildtype rescue transgene under the control of a ubiquitous promoter fully rescued embryonic muscle detachment of talin null embryos, introducing either the mutant ABS3 or the R2531G transgenes provided only a partial rescue of muscle detachment in the talin null background ([17]; Fig 8D–8F; Quantified in J). Furthermore, when we introduced the mutant ABS3 transgene into the background of the ABS2 mutant we significantly rescued the muscle attachment and recapitulated the transheterozygote phenotype (Fig 8H; quantified in 8J). These results validate our approach and confirm the specificity of the synergistic interaction between ABS2 and ABS3 talin mutants. In contrast, when the R2531G transgenes were introduced into the background of ABS2 mutant embryos we did not observe the strong rescue that was seen with the ABS3 transgene but a mild rescue similar to the effect of introducing the R2531G transgene into talin null flies (Fig 8F and 8I; quantified in 8J). These results suggest that the ability of talin to dimerize is key for the synergistic interaction we observe between talin ABS2 and talin ABS3 mutant alleles in line with the idea that talin functions as a dimer (Fig 8K).

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Fig 8. ABS2/ABS3 transheterozygote interaction requires talin dimerization.

(A-I) Confocal images of stage 17 whole-mount embryos stained with muscle cytoskeleton marker MHC (white) and Talin (magenta) ubiquitously (ubi) expressing: no transgenes (A,D,G), Talin ABS3 mutant protein (B,E,H) and Talin R2531G protein (C,F,I) in Wt (rhea^+/+, A-C), rhea null (rhea^-/-, D-F) and ABS2 mutant (rhea^ABS2/ABS2, G-I) backgrounds. Scale bar: 50 μm. (J) Number of detached muscles per embryo. Floating bar chart representing minimum and maximum value per genotype (n>10). Mean value represented by a solid line. Lower-case letter (a-e) indicates a different statistical group (P<0.05) using one-way ANOVA, Dunnett’s post-hoc. (K) Schematic model, created with BioRender.com, illustrating the role of ABS2 and 3 in integrin mediated adhesion. Solid arrows indicate recruitment to the MTJs whereas dashed ones represent the lack of recruitment.

https://doi.org/10.1371/journal.pgen.1011224.g008

Molecular biology

The generation of the ABS2 and ABS3 mutants described here (Fig 1) are based on a modified version of the following protocols: (http://flycrispr.molbio.wisc.edu, [47]). For ABS2 the following target sequences were respectively used for S1 and S2: 5′-GTAAGGCAGCGGAGGAACTTCGG-3′ 5′-GGCTACTTCGGACTTGGTGCAGG-3′. The 5’ and 3’ donor arms (Fig 1D) were synthesized by Integrate DNA Technologies (IDT) and assembled by Gibson cloning in the pHD-ScarlessDsRed plasmid (gift from Kate O’Connor-Giles, Addgene plasmid # 64703; http://n2t.net/addgene:64703; RRID:Addgene_64703). pHD-ScarlessDsRed is a donor vector with a 3xP3-DsRed marker cassette flanked by PBac transposon ends allowing for easy identification of engineered lines. For ABS3 the following target sequence was used 5′-GCTTATTTCCACAGCCAAGCAGG-3′. The single-stranded oligonucleotides (ssODNs) was generated by IDT. All targeting gRNAs were cloned by annealing the corresponding target sequence oligonucleotides into the pU6-BbsI-chiRNA plasmid[47] via the BbsI restriction sites. For both mutants the CRISPR injection mix containing the double-stranded donor DNA (500 ng/μl) along with targeting plasmids (100 ng/μl) was sent to Bestgene Inc. for injection. ABS2 mutant transgenic flies were identified by eye color screening using the DsRed gene included in the donor DNA. This visible eye marker was then excised by crossing ABS2 mutants to P{Tub-PBac\T}2 flies expressing piggyBac transposase leaving behind a residual TATA sequence. As the selection cassette was inserted into a native TATA site that was deleted in the donor vector the cassette was seamlessly removed. ABS3 mutant transgenic flies were identified by crossing P0 flies (n = 10) to third chromosome balancers then letting F1 flies interbreed. Resulting F2 progeny were crossed to rhea null flies for complementation test (100 single F2 flies crosses per P0 line). F2 candidates unable to complement rhea null were then sequenced for the presence of the ABS3 mutation. The resulting mutants were then recombined in an FRT2A background and sequenced using standard techniques (Fig 1F and 1G).

For the quantitative real-time PCR (qPCR) total RNA was isolated from whole flies using TRIzol. A total of 0.5 μg total RNA was converted into cDNA using the qScript cDNA synthesis kit (Quanta Biosciences). Subsequently, qPCR was performed using iQ SYBR Green Supermix (BIO-RAD). Talin mRNA levels were averaged between three independent experiments performed four times and normalized to β-tubulin expression. Primers used for Talin were located 3′ to the K17E mutation and were as follow 5′-GCCAGAACAATACTTTGGGTCG-3′ and 5′-AACTGGGCATTTCGCTGGAA-3′. β-tubulin expression was determined using the primer pair 5′-ATCATCACACACGGACAGGA-3′ and 5′-GAGCTGGATGATGGGGAGTA-3′.

Fly stocks and genetics

Except for viability/survival, motility and vinculin recruitment all experiments were performed in rhea mutant background such that wild-type maternal contributions of Talin were eliminated using the following alleles: rhea^79a (null allele, [15], rhea^ABS2, rhea^ABS3 or rhea^K17E [43] in conjunction with the dominant female sterile technique [74]. Females of the genotype yw, hs-Flp/+;; rhea mutant, FRT2A/OvoD1, FRT2A were subjected to a heatshock regime during the larval stages to generate mosaic animals in order to give rise to rhea mutant oocytes. Mosaic virgins were assessed for wing blisters and then crossed to the corresponding rhea mutant/TM6b, dfd-GMR-nvYFP males. Embryos without the fluorescent balancer were selected for analyses. ABS2 and 3 transheterozygotes mutants (rhea^ABS2/ABS3) were generated by crossing heatshocked yw, hs-Flp/+;; rhea^ABS2, FRT2A/OvoD1, FRT2A virgins to rhea^ABS3/ TM6b, dfd-GMR-nvYFP males. Full-length Vinculin with a C-terminal TagRFP tag [63] were driven in the muscles using the mef2-GAL4 driver. Vinculin::GFP allele was a gift from Yohanns Bellaïche [64]. GFP-tagged full-length talin construct (Talin-GFP, [75]) as well as Talin ABS3 (Talin KVK-DDD in [17]) and Talin R2531G [17] where ubiquitously expressed via the Ubiquitin-E63 gene promoter [75]. The ubi::TalinGFP line [47] used in rescue experiments has been shown to phenotypically rescue null rhea alleles fully to the end of larval stages but for currently unknown reasons fails to rescue rhea null mutants to viability.

Viability and motility assay

5 mm black dots were drawn at the back of 60x15 mm 2% agar petri dishes. 12–24 hours old eggs were then placed on each dot and monitored for 5 days. Resulting larvae were assessed for viability: most advanced developmental stage reached by at least ten larvae, L1 survival rate: percent eggs of a given phenotype hatching and L1 motility: percent L1s moving outside of 5 mm black dot after 1 hour minimum.

Confocal immunofluorescence imaging and image analysis

For all antibody stainings embryos were collected for 24 h at 25°C and heat fixed [48]. For rhodamine phalloidin staining, embryos were fixed in 4% formaldehyde followed by devitellinization in 80% ethanol. Antibody stainings were carried out according to standard procedures. Antibodies used were against the following proteins: MHC (mouse, 1:200; from Dan Kiehart, Duke University, Durham, NC), PINCH (rabbit, 1:1000; from Mary Beckerle, Huntsman Cancer Institute, UT), Talin (rabbit, 1:500), αPS2 (rat, 1:100; 7A10), Paxillin (rabbit, 1:1000) (Yagi et al., 2001)[56], tiggrin (mouse, 1:500; from Liselotte Fessler, UCLA, CA) and GFP (rabbit, 1:1000; A6455, Invitrogen). Fluorescently conjugated Alexa Fluor 488, Cy3 and Cy5 secondary antibodies were used at 1:400 dilution (Molecular Probes). Images were collected using an Olympus FV1000 inverted confocal microscope. For all micrographs of whole embryos, larvae or of MTJs, z-stacks were assembled from 8–12 0.5 μm confocal sections. Statistically significant differences were assessed by two-tailed Student’s t-tests in all cases, except when we sought to compare between multiple constructs, where one-way ANOVA was used. Differences between 2 percentage groups was assessed by chi-square test. Statistical analysis was carried out using Prism4 software (GraphPad, La Jolla, CA). Relative protein localization per larva was determined by the average pixel intensity of 3 MTJs divided by the average pixel intensity of the corresponding background area: zone of equal size directly anterior to the MTJ (Raw data shown in S1 Data). For intensity traces across MTJs (Fig 6P), the ImageJ plot profile tool was used to determine the average signal intensity across the boxed area indicated on the images. Intensity curves were obtained from unprocessed greyscale images so that first the peak intensity of each channel across the area of interest was set as 100%. Each curve was then normalized to the average intensity measured outside of the MTJ. Aggregates and muscle defects were counted manually using ImageJ.

Muscle detachment analysis

For embryonic muscle detachment analysis, z-stack projections of whole mount stage 17 embryos of each genotype stained with α-MHC were selected. Total number of muscles that were clearly detached and had a round shape was scored manually using ImageJ cell counter function (Raw data shown in S1 Data). For larval muscle detachment analysis desired genotypes were crossed into a fluorescent Z-disc marker (α-actinin-GFP). Resulting larvae were then heat-fixed by submerging them into 65°C PBS for 1–2 s and mounted in halo carbon before immediately being imaged [76].

Recombinant protein expression and purification

For biochemical analysis, pET151 plasmids for fly talin R4 Wt (residues 932–1062), R4-3E (residues 932–1062, K941E/Q942E/Q949E R7R8), R7R8 Wt (residues 1367–1666), R7R8-3E (residues 1367–1666 K1510E/R1520E/K1532E), R13 Wt (residues 2304–2489) and R13-3D (residues 2304–2489 K2450D/V2451D/K2452D) were produced as codon optimized synthetic genes by GeneArt.

15N-labeled proteins were prepared by transforming talin constructs into BL21(DE3)* E. coli cells and grown in 10 mL of LB + ampicillin at 37°C overnight as starter cultures and then in 2M9 minimal medium containing 15N-ammonium chloride at 37°C until an OD600 of 0.7–0.8. The cells were induced with 1 mM IPTG, then the cultures were incubated overnight at 20°C. The following day, harvested cells pellets were resuspended in 20 mM Tris-HCl, pH 8, 500 mM NaCl, and 20 mM imidazole, and lysed by sonication. Protein purification was achieved by nickel affinity chromatography using a 5 ml HisTrap HP column (GE Healthcare). The eluted proteins were dialyzed into 20 mM Tris-HCl, pH 8, 50 mM NaCl with AcTEV protease (Invitrogen) to remove the His-tag. After overnight dialysis, the proteins were purified with a HiTrap Q HP cation (R7R8 and R13) or HiTrap S HP anion (R4) exchange column (GE Healthcare).

Circular Dichroism (CD) experiments

Protein samples at a concentration of 50 μm were placed in a quartz cuvette and far-UV spectra were collected between 260 and 200 nm wavelengths using a JASCO J-715 spectropolarimeter at 20 and 95°C.

NMR experiments

15N-labeled protein samples at 0.15 mM were placed in Shigemi NMR tubes containing 10% D2O and 2D 1H-15N HSQC spectra were collected at 298 K on Bruker Avance III 600 MHz NMR spectrometer equipped with CryoProbe.