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The Drosophila formin Fhos is a primary mediator of sarcomeric thin-filament array assembly

  1. Arkadi Shwartz
  2. Nagaraju Dhanyasi
  3. Eyal D Schejter  Is a corresponding author
  4. Ben-Zion Shilo  Is a corresponding author
  1. Weizmann Institute of Science, Israel
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Cite this article as: eLife 2016;5:e16540 doi: 10.7554/eLife.16540

Abstract

Actin-based thin filament arrays constitute a fundamental core component of muscle sarcomeres. We have used formation of the Drosophila indirect flight musculature for studying the assembly and maturation of thin-filament arrays in a skeletal muscle model system. Employing GFP-tagged actin monomer incorporation, we identify several distinct phases in the dynamic construction of thin-filament arrays. This sequence includes assembly of nascent arrays after an initial period of intensive microfilament synthesis, followed by array elongation, primarily from filament pointed-ends, radial growth of the arrays via recruitment of peripheral filaments and continuous barbed-end turnover. Using genetic approaches we have identified Fhos, the single Drosophila homolog of the FHOD sub-family of formins, as a primary and versatile mediator of IFM thin-filament organization. Localization of Fhos to the barbed-ends of the arrays, achieved via a novel N-terminal domain, appears to be a critical aspect of its sarcomeric roles.

https://doi.org/10.7554/eLife.16540.001

eLife digest

Muscles owe their ability to contract to structural units called sarcomeres, and a single muscle fiber can contain many thousands of these structures, aligned one next to the other. Each mature sarcomere is made up of precisely arranged and intertwined thin filaments of actin and thicker bundles of motor proteins, surrounded by other proteins. Sliding the motors along the filaments provides the force needed to contract the muscle. However, it was far from clear how sarcomeres, especially the arrays of thin-filaments, are assembled from scratch in developing muscles.

When the fruit fly Drosophila transforms from a larva into an adult, it needs to build muscles to move its newly forming wings. While smaller in size, these flight muscles closely resemble the skeletal muscles of animals with backbones, and therefore serve as a good model for muscle formation in general. New muscles require new sarcomeres too, and now Shwartz et al. have observed and monitored sarcomeres assembling in developing flight muscles of fruit flies, a process that takes about three days.

The analysis made use of genetically engineered flies in which the gene for a fluorescently labeled version of actin, the building block of the thin filaments, could be switched on at specific points in time. Looking at how these green-glowing proteins become incorporated into the growing sarcomere revealed that the assembly process involves four different phases. First, a large store of unorganized and newly-made thin filaments is generated for future use. These filaments are then assembled into rudimentary structures in which the filaments are roughly aligned. Once these core structures are formed, the existing filaments are elongated, while additional filaments are brought in to expand the structure further. Finally, actin proteins are continuously added and removed at the part of the sarcomere where the thin filaments are anchored.

Shwartz et al. went on to identify a protein termed Fhos as the chief player in the process. Fhos is a member of a family of proteins that are known to elongate and organize actin filaments in many different settings. Without Fhos, the thin-filament arrays cannot properly begin to assemble, and the subsequent steps of growth and expansion are blocked as well.

The next challenges will be to understand what guides the initial stages in the assembly of the thin-filament array, and how the coordination between assembly of actin filament arrays and motor proteins is executed. It will also be important to determine how sarcomeres are maintained throughout the life of the organism when defective actin filaments are replaced, and which proteins are responsible for carrying out this process.

https://doi.org/10.7554/eLife.16540.002

Introduction

Sarcomeres constitute the basic functional units of muscle fibers, endowing these large and specialized cells with their contractile capacity. Central to sarcomere function is the lattice-like organization of two filament systems: an actin-based thin-filament array, which provides a stiff backbone along which thick filaments, composed of myosin motor proteins, 'slide' in order to produce force and contractile motion (Squire, 1997). The spatial organization and efficient operation of this remarkable cellular machinery relies on a host of dedicated proteins and protein complexes, which act to regulate sarcomere size and streamline its activity, and to coordinate between the multiple sarcomeric units that comprise individual myofibrils (Clark et al., 2002; Ehler and Gautel, 2008; Gautel and Djinovic-Carugo, 2016).

Despite their fundamental significance, elucidation of the molecular mechanisms underlying assembly, maturation and maintenance of thin-filament arrays remains one of the major open issues in the study of sarcomere structure and function. While mechanisms relating to size definition and stability of the arrays have been extensively investigated (Fernandes and Schock, 2014; Meyer and Wright, 2013), other key aspects of microfilament array formation and dynamics, including determination of distinct phases of array maturation, the identity and regulation of elements mediating filament nucleation/elongation, and the processes governing incorporation of additional filaments into nascent arrays are not resolved (Ono, 2010).

Here we address these matters in the context of formation and development of the Drosophila indirect flight muscles (IFMs). These are the largest muscles of the adult fly, which power flight by regulated contraction of the thorax (Dickinson, 2006). A major subset of the IFMs, the dorso-longitudinal muscles (DLMs), closely resemble vertebrate skeletal muscles in both their developmental program and in their mature myofibrillar structure (Dutta and VijayRaghavan, 2006; Roy and VijayRaghavan, 1999), making them a particularly attractive model system, in which the powerful molecular genetic approaches available to study Drosophila development can be harnessed to investigate and elucidate general principles of myogenesis.

DLM formation initiates by fusion of hundreds of individual myoblasts to a set of larval muscles during the first 24-30 hours of pupal development (Fernandes et al., 1991). The subsequent ~80 hrs of myogenesis leading up to eclosion of the adult fly include formation and maturation of a parallel arrangement of myofibrils and assembly and growth of sarcomeric units within them (Reedy and Beall, 1993Weitkunat et al., 2014). This sequence of events takes place over a wide time window, providing an opportunity to temporally dissected and manipulate the coordinated processes giving rise to thin-filament array assembly and maturation.

IFM sarcomeres initiate as small, nascent structures, that grow considerably over the course of pupal development (Sparrow and Schock, 2009), reaching a final, uniform size of 3.4 µm in length and 1.5 µm in diameter. Spatial organization of the mature, evenly-spaced IFM sarcomeres closely mirrors that of striated vertebrate skeletal muscle (Reedy and Beall, 1993). Individual sarcomeric units are defined by Z-disc borders, which serve as anchoring sites for the barbed ends of the thin-filament arrays, and by a central, microfilament-free H-zone, bordered by the pointed-ends of neighboring thin-filament arrays. We utilized temporally-controlled expression of GFP-tagged actin monomers (Roper et al., 2005) to follow thin-filament array dynamics, and recognize key phases and distinct transitions of the arrays throughout sarcomerogenesis. In parallel we identified Fhos, the single Drosophila member of the conserved FHOD family of formin proteins (Schonichen and Geyer, 2010), as a major contributor to thin-filament array assembly and growth. An important aspect of Fhos involvement is its critical role in radial growth of the arrays, by mediating incorporation of new peripheral filaments to the nascent core structure. The elongation of thin filament arrays, shown to occur primarily from their pointed ends (Mardahl-Dumesnil and Fowler, 2001), is mediated by the WH2-domain actin regulator Sals protein. While the combined activities of Fhos and Sals can account for most aspects of thin-filament array growth and maturation during pupal stages, other elements are likely involved in additional processes that shape and maintain the arrays, such as continuous monomer exchange at the barbed-ends.

Results

Distinct modes of thin-filament array assembly and growth

Assembly of sarcomeric thin-filament arrays has been traditionally studied by monitoring global changes in sarcomere structure in fixed samples of muscle tissue. However, in order to decipher the underlying regulatory mechanisms and uncover the machineries that drive this process, it is essential to monitor the dynamic patterns of actin monomer incorporation into the growing sarcomere. We therefore followed the incorporation of GFP-tagged actin monomers into IFM sarcomeres, throughout the pupal stages of development, as a means of revealing the assembly and maturation of IFM sarcomeric thin-filament arrays in developing flies. Inducible UAS-based GFP-actin transgenes (Roper et al., 2005; Verkhusha et al., 1999) have proven to be reliable tools and have been used extensively to study microfilament localization and dynamics during Drosophila development (Fulga et al., 2007; Jacinto et al., 2000; Kaltschmidt et al., 2002; Perkins and Tanentzapf, 2014; Schottenfeld-Roames and Ghabrial, 2012). The actin isoform at chromosomal position 88F, one of six Drosophila actin genes, is specifically expressed during the formation of the pupal muscles, and represents the major actin isoform used in this tissue (Beall et al., 1989; Fyrberg et al., 1983). We therefore employed temporally-restricted induction protocols of UAS-GFP-actin88F (Roper et al., 2005), and compared the resulting GFP patterns to the outlines of phalloidin-stained sarcomeres, as a primary tool for following the dynamics of IFM thin-filament array development (Figure 1A).

Figure 1 with 1 supplement see all
Four distinct modes of GFP-actin monomer incorporation contribute to formation of IFM thin-filament arrays.

(A) Scheme of IFM development intervals used for unrestricted and temporally restricted expression of GFP-actin88F (B–E”) or GFP-actin5C (G–I). (B–B”) Induction of GFP-actin88F expression (green, gray) with mef2-Gal4 throughout fly development results in full monomer incorporation into the thin-filament arrays (phalloidin- blue, gray), as monitored in IFMs of young (1–3 days old) adults. Z-discs are indicated by anti-Zasp52 (red). The designations 'Z' and M' are used throughout to mark the Z-disc (array barbed-end) and H-zone/M line (array pointed end) regions of the sarcomere. (C–E’’) Incorporation patterns of GFP-actin88F (green, gray) following temporally restricted expression pulses using the mef2-GAL4 driver and the GAL80ts/TARGET system. Microfilaments are visualized with phalloidin (blue, gray). Z-discs are indicated by anti-Zasp52 (red). (C–C”) 0–30 hrs APF. Initial uniform incorporation. (D–D”) 30–45 hrs APF. 'Patched' incorporation of monomers. This mode occurs mainly at array ends (insets in D and D’), in proximity to the future Z-disc (Z, white arrow) or towards the opposite boundary of the nascent arrays (M, red arrow). (E–E’’) 50–90 hr APF. Monomer incorporation into a 'frame' generated by peripheral 'thickening' (white arrowhead in panel E inset) and pointed-end growth (M and red asterisk in panel E inset). Red arrowhead (panel E’ inset) points to an absence of incorporated monomers at the barbed-end boundary (Z) of the arrays. (F) Schematic representations of the incorporation process. Blue filaments denote previously incorporated ('old') actin, while green (88F) and orange (5C) marks monomers newly incorporated during the indicated pulse. An initial period (0–30 hr APF) of extensive actin polymerization and the establishment of nascent thin-filament arrays are followed by an interim period (30–45 hr APF) of 'patchy' incorporation during which individual, uniform sarcomeric units are defined. The second half of pupal development is devoted to array growth via pointed-end elongation and recruitment of circumferential filaments, as well as turnover at array barbed ends. (G–I) Incorporation patterns of GFP-actin5C (green) following temporally restricted expression pulses using the GAL80ts/TARGET system. Z-discs are indicated by anti-Zasp52 (red). Restricted expression 'windows' corresponded to 40–90 (G), 50–90 (H) and 70–90 (I) hrs APF. Insets show the GFP-actin5C incorporation patterns in single sarcomeres, in which the prominent Z-disc associated stripe is outlined (Z, Z-disc region; M, M-line region). (J) Quantification reveals a constant width of the GFP-actin5C incorporation stripe overlying the Z-disc region despite the different pulse durations, n = 200 (50 sarcomeres each from 4 different flies). Scale bars correspond to 5 μm in all main panels, 2 μm in the insets.

https://doi.org/10.7554/eLife.16540.003

Following continuous expression of GFP-actin88F, a close correspondence between the sarcomeric GFP-actin and phalloidin patterns is observed in IFMs isolated from adult flies (Figure 1B–B”). The normal size and appearance of the IFM sarcomeres indicates that the GFP-tagged actin does not block or interfere with the structuring and organization of the thin filament arrays throughout the process, and the overlap between the phalloidin and GFP-actin patterns demonstrates the reliability of this tool for monitoring thin-filament array assembly.

We now attempted to break down into stages the processes of IFM thin-filament array organization and growth, by restricting actin-GFP expression to defined 'time windows' during pupal development. Temporal control of expression was achieved by using the general muscle driver mef2-GAL4 (Ranganayakulu et al., 1996) in combination with the GAL80ts/TARGET system (McGuire et al., 2004). While the timing of induction of GFP-actin is readily controlled using this system, effective 'chase' is not possible due to the stability of the GFP-actin monomers. We first induced expression of GFP-actin88F at the onset of pupariation, and examined the nascent IFMs at 30 hr APF (after puparium formation) (Figure 1C–C”). Individual myofibrils are already apparent at this early stage, displaying a nearly homogeneous distribution of microfilaments and an irregular pattern of nascent Z-disc structures, revealed using the early Z-disc marker Zasp52 (Jani and Schock, 2007; Katzemich et al., 2013). A full correspondence between the GFP-actin88F and phalloidin patterns is observed, implying that the bulk of early IFM microfilaments are formed by extensive de novo filament polymerization.

A very different profile of actin monomer incorporation is observed when a 15 hr pulse of actin88F-GFP expression is provided at 30–45 hr APF, immediately following the initial period of extensive polymerization (Figure 1D–D”). The GFP-actin and microfilament distributions become markedly distinct from each other during this phase, in which the patterns of Zasp52, the M-line marker Obscurin (Burkart et al., 2007) and phalloidin are now indicative of repeating sarcomeric units (Figure 1D–D” and Figure 1—figure supplement 1B–E). In contrast to the early 'smeared' pattern that filled the myofibrils, actin-GFP is now restricted to discrete, isolated spots, the great majority of which (~80%) positioned at either or both the ends of the nascent arrays (insets Figure 1D,D’ and Figure 1—figure supplement 1B–F). This pattern suggests that the initial establishment and structuring of individual sarcomere scaffolds, achieved during this interim phase of pupal development, relies on organization of the existing microfilaments, produced earlier (between 0–30 hr APF). Utilization of newly produced monomers during this interim phase of assembly is limited, and primarily involves 'patchy' incorporation along and (mostly) at the ends of nascent thin-filament arrays, thereby contributing to the generation of uniformly-sized sarcomeres.

A subsequent pulse, between 50 and 90 hr APF, revealed yet a third pattern of actin incorporation (Figure 1E–E” and Figure 1—figure supplement 1G–H”). During this (final) phase of pupal development, sarcomere units grow noticeably in both length and width, and the characteristic striated pattern of alternating Z-discs and filament-free H zones becomes clearly evident. Newly added actin is observed to form a distinct 'frame' that surrounds a dark rectangular core (insets Figure 1E and E’). The core presumably corresponds to the initial thin-filament array assembled during earlier phases. Conversely, the frame-like structure is likely composed of two separate contributions of newly synthesized actin to the nascent core array: extension of the initial fibers at their 'pointed' (M-line associated) ends, and addition of complete new fibers at the circumference of the sarcomere. This picture coincides well with previous studies of global IFM sarcomere growth (Mardahl-Dumesnil and Fowler, 2001; Reedy and Beall, 1993). Subdividing this relatively large interval further, by initiating GFP-actin88F production at 60 hrs APF, generated similarly shaped 'frames' of monomer incorporation, but of smaller size (Figure 1—figure supplement 1H–J). These observations demonstrate that both aspects of actin incorporation- pointed end growth and peripheral thickening- continue throughout the entire period. Monitoring of actin monomer incorporation patterns reveals therefore a dynamic, multi-faceted timeline of IFM thin-filament array assembly during pupal development (Figure 1F).

An additional, prominent feature of the GFP-actin pattern following the 50–90 hr APF pulse was a dark stripe overlying the entire Z-disc (insets Figure 1E’), implying that actin88F-GFP was not incorporated at this position, in neither the core array nor in newly added filaments at the periphery (Figure 1E,E’’). This observation suggests that elongation and thickening of the array during the final 50–90 hr APF interval are accompanied by a third mode of monomer incorporation, at the barbed end of the arrays. Absence of GFP-actin88F incorporation at the Z-disc was previously noted by Roper et al (2005), who also demonstrated preferential localization of other GFP-actin isoforms at this site, implying a specific, possibly steric hindrance of GFP-actin88F incorporation. We therefore chose to use the GFP-tagged version of actin5C (Roper et al., 2005), a ubiquitous isoform, as a tool for monitoring monomer incorporation dynamics at barbed ends of the thin-filament array. Expression of GFP-actin5C during the 40–90 hr APF time-window resulted in a nearly complementary incorporation profile to the one generated by actin88F-GFP: a prominent stripe of GFP-actin adjacent to the Z-disc and relatively limited incorporation in the peripheral actin strands and pointed ends (Figure 1G). Initiation of the GFP-actin5C pulse at different times within this 50 hr interval, generated a thin bright incorporation stripe of constant width adjacent to the Z-disc in all cases (Figure 1H–J). This result implies continuous exchange and turnover of actin, rather than actual growth at the barbed ends of the arrays, and is consistent with the notion that lateral growth of array filaments occurs primarily at their pointed ends (Littlefield et al., 2001; Mardahl-Dumesnil and Fowler, 2001; Molnar et al., 2014).

The correspondence between the regular length of the sarcomeres and the size of the actin monomers can provide a rough estimate of the number of actin monomers that build the entire structure and the proportion undergoing turnover. In a mature sarcomere, the length of thin-filament actin fibers (as measured from the Z-disc to the H-zone) is ~1.70 µm. It is difficult to accurately measure the width of the domain of dynamic actin-monomer exchange due to the limitations of light microscopy resolution, but it is roughly 0.15–0.3 µm on each side of the Z-disc. Given an estimated actin subunit size of 2.7 nm (Sept et al., 1999), we can say that a complete thin filament is comprised of ~650 monomers, whereas the zone of continuous exchange at the Z-disc encompasses 50–100 monomers.

The formin protein Fhos is a major mediator of sarcomeric thin-filament array assembly and growth

To identify actin regulators that are involved in the different phases of IFM thin-filament array organization and growth, we focused on members of the formin protein family, which are major mediators of nucleation and elongation of linear microfilament arrays (Campellone and Welch, 2010). The Drosophila genome harbors six members of this protein family, each representing a distinct formin sub-family (Liu et al., 2010; Mi-Mi et al., 2012). To assess their involvement in the IFM sarcomere formation, we used the muscle-specific driver mef2-Gal4 to induce expression of RNAi directed against each of the six formins throughout development via UAS-based transgenic constructs, and examined IFM morphology following their isolation from newly eclosed or pharate adults. In most instances, IFM development was only mildly affected, if at all, following individual knockdown of the different Drosophila formins.

A severe IFM phenotype was obtained, however, following an expression of RNAi constructs directed against Fhos, the single Drosophila FHOD sub-family homolog (Schonichen and Geyer, 2010). A normally-sized set of six DLM fibers formed in Fhos knockdown flies (Figure 2—figure supplement 1A,B), indicating that the IFM developmental program is properly initiated. However, the internal organization of these fibers was severely disrupted. This was made apparent by staining the DLMs for key structural components, including α-actinin as a marker for sarcomeric Z-discs, microfilaments and muscle myosin (Figure 2A–B”). Fhos knockdown DLMs appear to contain myofibril-like elements, but these are thin and randomly oriented (Figure 2B). Furthermore, in contrast to the highly regular division of wildtype myofibrils into repetitive sarcomeric units (Figure 2A,A’), the abnormally thin Fhos knockdown myofibrils display only sporadic α-actinin -stained structures, and a 'smeared', uneven distribution of microfilaments (Figure 2B,B’). In addition, muscle-specific myosin is disorganized, and to a large extent lacks an obvious association with microfilaments (Figure 2B,B”).

Figure 2 with 2 supplements see all
The formin Fhos is essential for organization and growth of thin filament arrays.

(A–C”) Confocal images of IFMs dissected from 1 day old flies or pharate adults and stained with anti- α-actinin (red) to mark Z-disc structures, phalloidin (blue, gray) to visualize microfilaments and anti-MHC (green, gray) to visualize myosin. (A–A”) mef2-GAL4 control. Z and M mark the Z-disc and M-line of a single sarcomere. (B–B”) mef2-GAL4>UAS-fhos RNAi (knockdown of all fhos isoforms). Myofibril and sarcomere structure and organization are defective, but sporadic, undersized sarcomeric units can be observed (white arrowhead in B). (C–C’’) fhosΔ1/Df(3L)BSC612 (fhos null). Deletion of the fhos locus results in full impairment of myofibril and sarcomeric organization. (D–E) TEM micrographs of longitudinal sections of IFMs dissected from control 1 day old flies (D) and fhos null (fhosΔ1/Df(3L)BSC612) pharate adults (E). Distinction in the overall myofibril organization is readily apparent, with fhos null IFMs lacking typical myofibril and sarcomeric individualization. The insets contrast the stereotypic, highly-ordered structure of the control sarcomeric units (inset D) with the poor organization of arrays within fhos null myofibrils and their failure to form individual sarcomeres (inset E). Red arrowheads in (E) point to dispersed, rudimentary Z-discs. (F–G’) TEM micrographs of transverse sections of IFMs dissected from control 1 day old flies (F,F’) and fhos null (fhosΔ1/Df(3L)BSC612) pharate adults (G,G’). Primed panels are magnifications of the dashed squares in panels (F) and (G). In contrast to the highly ordered hexagonal lattice of thick (orange) and thin filaments (blue) in control myofibrils (F’), fhos null myobrils lack a defined spatial organization (G’). Scale bars: 5 μm (A-E), 500 nm (insets in D,E, and F,G), 100 nm (F’,G’).

https://doi.org/10.7554/eLife.16540.005

We sought to complement and enhance the analysis of Fhos-knockdown IFMs by studying mutant alleles in the Fhos locus. A recent study (Lammel et al., 2014) described several such alleles, including FhosΔ1, a small deficiency that completely removes the coding regions of most Fhos isoforms, and thus represents a severe, possibly null, gene disruption. FhosΔ1 homozygotes die as pharate adults, allowing to assess the effects of Fhos gene knockout on IFM development. Immuno-fluorescent staining with informative markers revealed that in FhosΔ1 hemizygous flies that reach the pharate adult stage, DLMs are highly disorganized, lacking even the trace appearance of sarcomeric units observed in Fhos knockdown DLMs (Figure 2C–C”).

We extended this study by subjecting the mutant DLMs to transmission electron microscopy (TEM) analysis. Longitudinal TEM sections underscored the disorganized nature of the Fhos mutant DLMs, which appear to be composed of irregularly shaped myofibrils, lacking a defined spatial orientation (Figure 2D,E). While myofilament arrays can be found within these structures, they fail to exhibit any of the features of regularly spaced sarcomeric units characteristic of wildtype DLMs (Figure 2D), and display only a few sporadic electron-dense spots that may represent rudimentary Z bands (Figure 2E). In contrast to the highly-ordered hexagonal lattice of thick and thin filaments within wildtype myofibrils, revealed by TEM cross-sectional views (Figure 2F,F’), the myofilament arrays in Fhos mutant DLMs are small, irregularly-spaced, and lack a defined spatial organization (Figure 2G,G’).

Highly defective myofibril and sarcomere organizations were already clearly apparent in FhosΔ1 hemizygotes at 50 hrs APF via both light microscopy (Figure 2—figure supplement 1E–F’) and TEM Figure 2—figure supplement 1G–J) analyses, demonstrating that the mutant phenotypes are a consequence of developmental abnormalities initiating at the onset of IFM sarcomere formation, rather than deterioration of normally formed structures. The severe phenotypes of Fhos knockdown and null mutant pupae demonstrate an essential role for Fhos in the assembly and organization of sarcomeric units within IFMs. The nearly complete lack of sarcomeric organization within myofibrils in the absence of Fhos activity implies a critical requirement for Fhos already at early stages of sarcomere assembly.

Such early arrest in sarcomerogenesis may mask potential requirements for Fhos at later stages of the process. To address this issue, we induced RNAi directed at Fhos using the IFM-specific act88F-Gal4 driver (Gajewski and Schulz, 2010), thereby delaying onset of Fhos knockdown to a more advanced phase of IFM development. While such flies emerged from the pupal case, they were flightless. A more detailed examination following isolation of IFMs revealed the establishment of an ordered array of intact, regularly-spaced sarcomeric units (Figure 2—figure supplement 2A–B’). However, these Fhos knockdown sarcomeres exhibited significantly shorter widths and lengths than sarcomeres from age-matched controls (Figure 2—figure supplement 2C,D), implying a requirement for Fhos in the elongation and peripheral thickening mechanisms underlying thin-filament array maturation.

Taken together, the range of phenotypic abnormalities associated with the various forms of disruption to Fhos function suggest that Fhos is a major mediator of sarcomere formation, contributing throughout pupariation to different aspects of thin-filament array assembly and maturation.

Localization of Fhos to Z-discs is critical for function

The Fhos locus is composed of two classes of transcripts, utilizing different promoters (Figure 3A). Both transcript classes share a set of 3’ exons, but differ in their 5’ regions, which include distinct non-coding and coding exons. As a result, the locus generates two main Fhos protein isoforms, in which a conventional FHOD-family formin, containing all of the canonical formin regulatory and actin-related functional domains, is appended to different N-terminal segments (Figure 3B). The isoform encoded by transcripts RA-RG features a short (63 residue) N-terminal domain, while transcripts RH-RJ encode an isoform bearing distinct and substantially larger N-terminal region that is conserved among Neopteran winged-insects (Bechtold et al., 2014).

Figure 3 with 1 supplement see all
Localization of Fhos at the Z-disc is essential for its function.

(A) Map of the Fhos genomic locus and transcripts (after Flybase, (Attrill et al., 2016). Shown are the nine known fhos transcripts (designated Fhos-RA--Fhos-RI), which are divided into two groups (RA-RG and RH-RI). The two groups share a nearly identical set of 3’ exons (red dashed rectangle), which encode a conventional FHOD-family formin, but are expressed via distinct regulatory regions, and possess distinct sets of 5’ exons, including 5’ coding exons (blue and purple dashed rectangles) that encode different N-terminal domains. The two transcript variants, RH and RA, used to generate, respectively, the long and short transgenic UAS-Fhos constructs are indicated by orange arrows. The insertion positions of three MiMIC elements, MI04231 (inserted downstream of long isoform initiation sites), MI01421 (inserted downstream of all transcript initiation sites), and MI09324 (used to produce the Fhos-GFP 'protein trap') are indicated by inverted triangles. Positions of two dsRNA target sequences used, one common to all fhos isoforms (red bar) and the other specific to the long forms (blue bar) are shown above the transcript map. The CRISPR/Cas9-generated deletion of the guanine residue at position 99 of the short isoform transcript and its adjacent sequence are indicated. (B) Schematic representation of three representative Fhos protein isoforms. The canonical formin domains common to all forms are colored red, while the alternative N-terminal domains are in blue (long forms) and purple (short form). Canonical domains indicated include the GTPase binding domain (GBD), formin homology (FH) domains 1/2 and 3, and the diaphanous autoregulatory domain (DAD). The positions of the I966A point mutation in the FH2 domain and the premature stop codon, generate by the frameshift mutation ΔG99 in the Fhos-PA N-terminal domain are indicated. (C–D’) Zasp (red) and phalloidin (blue and gray) stainings demonstrate the severe, null-like disruption of myofibril and sarcomere microfilament organization in hemizygous FhosMI01421/Df(3L)BSC612 pharate adult flies (C,C’), similar to that observed in fhosΔ1 hemizygotes. No rescue is observed following expression of UAS-GFP-Fhos-PA (green) driven by arm-Gal4 in this background (D,D’). (E–F) α-actinin (red) and phalloidin (blue and gray) stainings demonstrate the severe, null-like phenotypes following specific RNAi mediated knockdown of the Fhos long-isoforms (E,E’) and in hemizygous FhosMI04231/Df(3L)BSC612 (F) pharate adult flies. (G) Zasp (red) and phalloidin (blue) stainings demonstrate normal myofibril and sarcomeric structure of Fhos ΔG99/Df(3L)BSC612 hemizygotes, in which the short Fhos isoforms are not expressed. (H–L’’) Fhos localization in myofibrils, as monitored at two distinct pupal developmental time points, 45 hr APF (H–I’), and 65 hr APF (J–J’’), via a GFP 'exon trap' engineered at the insertion site of the MiMIC transposon MI09324 (green triangle in A). The GFP-tagged Fhos proteins (all isoforms) generated in this manner are visualized with anti-GFP (green or gray), Z-discs are visualized with anti-α-actinin or anti-Zasp (red), thin filament pointed ends visualized by anti-Tmod (blue) and microfilaments with phalloidin (blue). The diffuse/punctate initial localization of Fhos-GFP overlying broad portions of the growing myofibrils (H), in some cases shows an adjacent localization to the nascent Z-disc (I white arrowhead) or to array pointed ends (I’ red arrowhead). The initial punctate localization gives way to a striated pattern restricted to the vicinities of both the barbed (Z) and pointed (M) ends of the thin-filament arrays (J–J’’). (K–K”) Localization of the short isoform of Fhos in IFMs from a young adult fly, visualized by expression of UAS-GFP-Fhos-PA using the mef2-GAL4 driver (anti-GFP, green or gray). Z-discs are visualized with anti-α-actinin (red), and microfilaments with phalloidin (blue). GFP-Fhos-PA localizes to the vicinity of the pointed ends of the arrays (M). (L–L”) Localization of the long isoforms of Fhos in IFMs from a young adult fly, visualized with anti-HA (green and gray), following expression of UAS-HA-Fhos-PH using the mef2-GAL4 driver. Fhos-PH-PA localizes to the vicinity of the barbed ends of the arrays (Z), where it overlaps with the general Fhos distribution to both the barbed and pointed ends (M) of the arrays (visualized with anti-Fhos [red]). Microfilaments visualized with phalloidin (blue). Scale bars in all panels correspond to 5 μm.

https://doi.org/10.7554/eLife.16540.008

The short Fhos variant, represented by form RA, has been shown to rescue Fhos null mutant flies to adult viability, and to restore normal function in affected tissues (e.g. macrophage motility and wing inflation), when expressed ubiquitously via armadillo-GAL4 (Lammel et al., 2014). We were therefore surprised to discover that the flight muscles of such EGFP-Fhos-PA-rescued flies continued to exhibit severe Fhos mutant phenotypes (Figure 3C–D’), implying that the short form of Fhos, which mediates most Fhos developmental functions, is insufficient in this context. These observations raised the possibility that the larger isoforms, which have not been extensively studied, provide Fhos activities necessary for IFM sarcomere formation.

To address this issue directly, we utilized a transgenic RNAi construct specifically targeting the large 5’ coding exon (Figure 3A). Expression of this RNAi construct in muscle cells, which should eliminate only the long isoforms in this tissue, led to a strong disruption of sarcomere organization, closely resembling null mutations (Figure 3E,E’; see also [Schnorrer et al., 2010]). Furthermore, we observed similar deleterious effects on the IFM organization in flies hemizygous for MI04231 (Figure 3F), a MiMIC insertion allele that is predicted to specifically disrupt the long Fhos isoforms and leave the short isoforms intact (Figure 3A). These results lead us to conclude that the long N-terminal protein domain is critical for IFM function of Fhos.

To determine whether the long Fhos isoforms are sufficient, we used a CRISPR/Cas9 approach to generate small deletions in the exon encoding the N-terminal region of the short Fhos isoforms. One of these, FhosΔG99, results in a single nucleotide deletion, generating a translational frameshift and a predicted termination of translation of the short Fhos isoform after only 58 residues (Figure 3A,B). FhosΔG99 hemizygous flies, which do not express functional short Fhos isoforms, are fully viable and fertile, and do not exhibit any obvious morphological defects. Importantly, the IFMs of these flies display normal myofibril and sarcomere organization (Figure 3G). These observations imply that the long Fhos isoforms are sufficient for proper IFM myogenesis, and furthermore, that they can provide most, if not all, functional requirements for Fhos.

Identifiable protein domains are not found within the 1198 residue long protein sequence encoded by the large 5’ exon, which raised the possibility that this extension to the canonical FHOD-like formin provides a localization cue, rather than an additional functional moiety. To pursue this notion, we set out to determine the localization patterns of the different Fhos isoforms. We first made use of a MiMIC transposable element insertion in the Fhos gene locus (MI09324) and the RMCE technique (Venken et al., 2011) to generate a GFP 'protein trap' (Fhos-GFP), so that all isoforms of endogenous Fhos would also harbor a GFP tag (Figure 3A). Monitoring the Fhos-GFP signal at 45 hr APF revealed an initial diffuse localization to myofibrils, with some enrichment over nascent sarcomeric units (Figure 3H–I’). At later stages (65 hr APF), the localization of Fhos refines to discrete stripes overlying the barbed and pointed ends of the microfilament arrays within the sarcomere (Figure 3J–J’’), a pattern that persists throughout pupal stages and is still observed in young adult flies (Figure 3—figure supplement 1A–A”’). Staining with an antibody we raised to the Fhos C-terminal domains shared by all isoforms of the protein, displayed a similar pattern (Figure 3—figure supplement 1A–A”’).

We next examined the localization patterns of representative tagged versions of the short and long isoforms of Fhos, following their expression in IFMs. EGFP-Fhos-PA, representing the shorter isoform, was found to localize exclusively to the vicinity of sarcomere M-lines, corresponding to the pointed-ends of the thin-filament arrays (Figure 3K–K”). This observation raised the possibility that the short isoform is not functional on its own in IFMs due to its restricted localization pattern, which does not include the Z-disc associated barbed-ends of the thin-filament arrays. Notably, instances of ectopic localization of EGFP-Fhos-PA to Z-discs, which were sporadically observed when this construct was over-expressed in a null Fhos mutant background, were associated with markedly improved organization of the affected sarcomeres (Figure 3—figure supplement 1B–B”’). This observation supports the notion that localization of Fhos to the Z-disc region is critical for its function in the growing sarcomere.

To monitor localization of the long Fhos isoform, we generated an HA-tagged version of Fhos-PH, which contains 410 residues of the novel N-terminal domain (Figure 3B). Remarkably, HA-Fhos-PH was found to localize exclusively to the Z-disc region of IFM sarcomeres following expression via mef2-GAL4 (Figure 3L–L”), in complementary fashion to EGFP-Fhos-PA. Taken together with the genetic analysis, which identified the long isoform as the functional Fhos variant in IFMs, we conclude that localization of Fhos to the Z-disc/array barbed-end region, mediated by the long, novel N-terminal domain, is critical for its sarcomeric function. Interestingly, despite the localization of the HA-Fhos-PH construct to the Z-disc region, it only partially rescued the sarcomere organization defects of Fhos null flies (Figure 3—figure supplement 1C–C”’). In addition, we observe that the long Fhos isoforms, which provide full sarcomeric functionality, localize to both the Z-disc and M-line regions in FhosΔG99 hemizygotes (Figure 3—figure supplement 1D). Thus, while Z-disc localization of Fhos is an essential requirement for proper sarcomere assembly, we cannot rule out that Fhos performs functional roles at additional sites within maturing IFM sarcomeres.

The roles of Fhos in actin incorporation into thin-filament arrays

Having established that Fhos is a major contributor to IFM sarcomere organization, we now sought to elucidate its specific roles, by monitoring actin monomer incorporation patterns in the absence of Fhos function. Towards this end, we examined IFMs from Fhos knockdown pupae, following a restricted expression of GFP-actin88F during early (0–30 hr APF), interim (30–60 hr APF) and late (60–90 hr APF) phases of pupal development (Figure 4A). During the initial stages of IFM development, incorporation of GFP-actin88F into unstructured microfilament arrays within nascent myofibrils proceeded normally in Fhos knockdown IFMs (Figure 4B,C). This finding implies that Fhos is not essential for the initial 'burst' of strong polymerization activity characteristic of this phase (Figure 1C–C”), and is consistent with the establishment of properly sized but internally disorganized DLM myofibers in Fhos knockdown and mutant flies (Figure 2, Figure 2—figure supplement 1).

Fhos is required for the 'patchy' actin monomer incorporation and radial expansion aspects of thin-filament array growth.

(A) Scheme of IFM development intervals used for temporally restricted expression of GFP-actin88F (B–J) or GFP-actin5C (K–M’) in wildtype (B–B”,D,D’,H,H’,K,K’) or Fhos knockdown (C–C”,F,F’,I,I’,L,L’) IFMs. (B-I’) GFP-actin88F (green, gray) expression between 0–30 (BC’), 30–60 (D-G) and 50–90 (H-I’) hr APF. Z-discs are visualized with anti-Zasp52 (red) and microfilaments with phalloidin (blue, gray). (B–C’) The general and uniform incorporation of monomers into microfilaments characteristic of the initial phase of sarcomere formation (B–B”) is not affected in fhos knockdown myofibrils (C–C”). (D–G) The dispersed and 'spotty' wildtype incorporation pattern during the interim (30–60 hr APF) phase (D) is replaced by a pointed-end centered pattern in fhos knockdown myofibrils (F). These pattern distinctions are further demonstrated by heat maps of the GFP-actin88F distribution (D’,F’) and quantification of GFP intensity (E,G) derived from 10 μm profiles covering approximately four sarcomeric units (white lines in D and F; data were acquired for 15 profiles from 7 different pupae for each genotype [n = 105]). (H–I’) The incorporation 'frames'normally generated by late GFP-actin88F expression pulses (H,H’) lack peripheral incorporation (arrowheads) following fhos knockdown (I,I’), but these abnormally thin myofibrils retain proper incorporation at array 'pointed' ends (red asterisks). The lack of incorporation 'frames' is also demonstrated by heat maps (H’,I’), and the quantification of GFP intensity (J; data were acquired for 100 profiles from 4 different flies for each genotype [n = 400]) along vertical profiles (white dashed line H’,I’), which show the loss of peripheral incorporation and thinner sarcomeres (red dashed lines in J). (K-M) GFP-actin5C (green, gray) expression between 50–90 hr APF. Z-discs are visualized with anti-Zasp52 (red) and microfilaments with phalloidin (blue). GFP-actin5C induction in parallel to fhos knockdown showed normal Z-disc associated turnover (arrowheads). (M) Quantification of Z-disc incorporation band width (data were acquired for 50 Z-discs from 4 different flies for each genotype [n = 200]). Scale bars in all panels correspond to 5 μm.

https://doi.org/10.7554/eLife.16540.010

In contrast to these observations, a marked effect of Fhos knockdown on the actin monomer incorporation pattern can be discerned during the interim (30–60 hr APF) period of pupal development (Figure 4D–G). While wildtype IFMs display an irregular 'patchy' pattern of incorporation spread out over the nascent sarcomeric arrays (Figure 4D–E), Fhos knockdown IFMs exhibited a repetitive, undulating pattern, with peaks of incorporation centered at the pointed-ends of the arrays (Figure 4F–G). Fhos therefore appears to mediate monomer incorporation into filament patches, but is not involved in the emerging process of array elongation from pointed ends. This feature of Fhos knockdown IFMs persists during the final phase of pupal development, when wildtype sarcomeres display a frame-like pattern of incorporation (Figure 4H–J). While such 50–90 hr old Fhos knockdown IFMs retain pointed-end incorporation, they appear to be thinner and lack the circumferential accumulation of incorporated GFP-actin88F, which represents radial growth of the arrays through addition of peripheral microfilaments (Figure 4I,J), implying a requirement for Fhos in the array 'thickening' process. Finally, a GFP-actin5C incorporation band of normal width was readily detected adjacent to sarcomere Z-discs in Fhos knockdown IFMs (Figure 4K–M), implying that actin monomer exchange at barbed ends of the arrays is Fhos independent.

Analysis of actin monomer incorporation patterns thus provides a higher resolution and reveals specific roles for Fhos in mediating thin-filament array assembly and growth. These include the organization of microfilaments into nascent structures, shaping sarcomeres into uniformly sized and regularly-spaced units and, finally, radial growth of the arrays via peripheral thickening. On the other hand, the synthesis of the initial pool of microfilaments, array elongation from pointed ends and actin exchange at the Z-disc do not require Fhos, and thus rely on the activity of other actin regulators.

Sals is required for thin filament elongation from the pointed ends of the array

As the incorporation data suggests that filament extension from the pointed-ends of the arrays does not require Fhos, we sought to identify alternative elements that may mediate this key aspect of sarcomere growth. Such factors are unlikely to include formin family members, since formins are primarily thought to extend actin filaments from their barbed ends (Campellone and Welch, 2010; Goode and Eck, 2007). The WH2-domain protein Sarcomere length short (Sals) is an attractive candidate, since it was shown to contribute to pointed-end filament elongation in Drosophila larval muscles, and to be localized to the pointed ends of thin-filament arrays in IFM sarcomeres (Bai et al., 2007).

We first assessed the role of Sals in IFM sarcomerogenesis by using the mef2-GAL4 driver and the GAL80ts/TARGET system to express an RNAi construct targeted against sals from the onset of pupariation, and examined the effect on IFMs of pharate adult flies. The overall length of the sals-knockdown thin-filament arrays is significantly shorter than control arrays, and their pointed end borders are abnormally shaped and discontinuous (Figure 5A–C’, Figure 5—figure supplement 1D), consistent with a pointed-end extension function for Sals.

Figure 5 with 1 supplement see all
Sals is required for 'pointed-end' thin-filament growth.

(A) Scheme of IFM development intervals used for temporally restricted expression of sals RNAi and GFP-actin88F. (B–C’) IFMs dissected from control (mef2-GAL4) 1 day old flies (B,B’) and sals knockdown pharate adults (C,C’), in which RNAi expression was initiated at 0 hr APF. Z-discs are visualized with anti-Zasp52 (red) and microfilaments with phalloidin (blue or gray). sals knockdown results in sarcomere shortening and 'pointed' end abnormalities (insets in B’ and C’; for quantification see Figure 5—figure supplement 1D). (D–E’) IFMs dissected from young (1–2 day old) flies in which GFP-actin88F expression (anti-GFP, green, gray) was initiated at 50 hr APF on its own (D,D’) or together with sals RNAi (E,E’). M lines are visualized with anti-Obscurin (red) and microfilaments with phalloidin (blue). (F) The GFP-actin88F incorporation band at the 'pointed' ends is significantly decreased following sals knockdown, as shown by the M/Z intensity ratio (p<0.0001, P values determined by Mann-Whitney test), while the addition of peripheral microfilaments is unaffected. (G) Quantification of phalloidin intensities derived from 13 μm profiles (white line in D). sals RNAi myofibrils (red line) exhibit shorter sarcomeric units compared to control (blue line). Scale bars in all panels correspond to 5 μm.

https://doi.org/10.7554/eLife.16540.011

We next utilized the GFP-actin88F incorporation assay to further examine Sals activity in developing IFMs. Specifically, sals function was disrupted by knockdown beginning at 50 hr APF, when 'core' thin-filament arrays have already formed, monomer incorporation was monitored in young adults. The resulting incorporation pattern is a near 'mirror-image' of Fhos knockdown during this period: shortened sarcomeres displaying normal peripheral thickening of the core arrays (Figure 5D–E’,G), coupled with a significant decrease of actin incorporation at their pointed ends (Figure 5F).

The conserved pointed-end capping protein Tropomodulin (Tmod) (Gregorio and Fowler, 1996) is a second factor that could contribute to filament pointed-end elongation. Induction of RNAi targeting tmod at 0 hr APF indeed results in a significant shortening of IFM thin-filament arrays, but to a considerably lesser extent than the shortening observed following knockdown of sals (Figure 5—figure supplement 1A–D). Furthermore, tmod knockdown during the second half of pupal development only weakly affects the array length and pointed-end incorporation of actin monomers (Figure 5—figure supplement 1E–H).

Taken together, these results identify Sals as a major actin regulator, specifically mediating pointed-end growth of thin-filament arrays throughout IFM maturation, while the contribution of Tmod appears to be restricted to early stages of IFM development.

The barbed-end associated capacities of Fhos are dispensable during the initial stages of sarcomere organization

Formins employ a variety of molecular mechanisms for regulating microfilament organization and dynamics, including microfilament nucleation, elongation, bundling and capping activities (Goode and Eck, 2007; Harris and Higgs, 2006; Schonichen and Geyer, 2010). To begin to address this issue in the context of Fhos IFM function, we used CRISPR/Cas9 technology to insert a point mutation into the endogenous Fhos locus, thereby generating a single amino acid substitution (I966A according to residue numbering of the short form of Fhos, Figure 3B). Mutating this highly conserved residue in a variety of formins, including the mammalian Fhos homolog FHOD3, consistently abolished activities requiring barbed-end association (actin nucleation, elongation and capping) (Harris et al., 2006; Taniguchi et al., 2009; Xu et al., 2004).

FhosI966A hemizygous flies are viable and exhibit an externally normal morphology, implying that the nucleation activity of Fhos is generally dispensable. However, the IFMs of these flies contain abnormally thin myofibrils. Most of these myofibrils display, nevertheless, an organized structure of repeated sarcomeric units, with clear demarcation of the Z-discs (Figure 6A–A”), suggesting an arrest in sarcomere growth following proper initial assembly and organization. This notion was further borne out following TEM-level visualization, which showed that the thin FhosI966A myofibrils house properly structured and evenly-spaced sarcomeres (Figure 6B), harboring well-ordered lattices of thick and thin filaments (Figure 6C).

Early Fhos function does not require barbed-end activities.

(A–A’’) IFM myofibrils from a FhosI966A/Df(3L)BSC612 1 day old adult fly. Z-discs are marked by anti-α-actinin (red or gray) and microfilaments are visualized with phalloidin (blue or gray). The thin myofibrils display organized arrays of repeated sarcomeric units (red outlines in A’’). (B–F) TEM analysis of IFM myofibrils from FhosI966A/Df(3L)BSC612 flies. (B,C) One day old adult flies. A longitudinal section (B) shows a stereotypic sarcomeric unit displaying clear Z-disc (Z) and M line (M) structures. A cross section (C) shows an individual sarcomeric unit (red dashed circle) harboring a well-formed lattice of thick and thin filaments. Arrows point to accumulations of nearby filaments, which could serve as a source for radial growth, but have not been recruited. The degree of lattice organization within and outside the sarcomere can be appreciated from the spatial arrangement of representative thick (orange) and thin (blue) filaments. (D) Quantification of sarcomere size in wildtype and FhosI966A/Df(3L)BSC612 pupae and adult flies at the indicated ages, based on the number of thick filament units in TEM cross-sections (n = 20 for each background). (E,F) 50 hr APF pupae. Longitudinal (E) and cross (F) sections show myofibril individualization (red dashed lines in E) and formation of nascent sarcomeric units (red dashed circles in F) with defined Z-disc borders (red arrowheads in E; see also Figure 2—figure supplement 1G,H). Scale bars correspond to 5 μm (A), 100 nm (B), 500 nm (C,F) and 200 nm (E).

https://doi.org/10.7554/eLife.16540.013

The actual radial size of FhosI966A sarcomeres was assessed by determining the number of thick filament units in myofibril TEM cross sections. This analysis revealed that the sarcomeres of 1 day old Fhos-I966A mutants are ~ 8 times smaller than wildtype sarcomeres from similarly aged flies, and correspond in size to normal sarcomeres between 50–60 hr APF (Figure 6D). This observation is in keeping with the notion of an arrest in sarcomere growth during intermediate pupal stages. Furthermore, the appearance and organization of sarcomeres within myofibrils of FhosI966A hemizygotes at 50 hr APF (Figure 6E,F) closely matches those of sarcomeres from age-matched wildtype pupae (Figure 2—figure supplement 1G,H).

These light and electron microscopy analyses suggest therefore that the normal involvement of Fhos at the initial stages of IFM sarcomere assembly does not require a barbed end-associated activity, consistent with the early diffuse localization pattern of Fhos in myofibrils. However, the barbed-end localization and associated activities of Fhos appear to be essential for the maturation of nascent arrays, implying a multi-faceted involvement of this formin in sarcomere formation.

Discussion

Formation of the adult Drosophila IFMs during pupariation provides an established model system to study formation of skeletal muscles, and in particular the generation of the repeated sarcomere structure, the core functional unit underlying muscle contractility. The IFM system possesses several key features that make it amenable for detailed analysis. These include an extended developmental time window of ~60 hr, the availability of genetic methods for investigation (e.g., RNAi-based knockdown) and the highly ordered and repetitive organization of sarcomeric units within IFM myofibrils, which allows for the detection and analysis of both major and subtle alterations and defects. Our study focused on the actin based thin-filament array component of sarcomeres. Towards this end, we utilized an additional, highly useful feature of the IFM system: the capacity to monitor the incorporation pattern of new actin monomers at discrete stages of the process, using transgenic GFP-actin constructs whose expression can be readily manipulated. The analyses performed using the various approaches and tools available for the study of the IFMs allow us to chart the principles, timeline and molecular basis for assembly, organization and maturation of thin-filament arrays within this model of sarcomerogenesis.

While it is likely that regulators of linear actin belonging to the formin family play a central role in the formation and maintenance of actin filament arrays in the sarcomere, redundancy among them appears to be commonplace in this context (Mi-Mi et al., 2012; Rosado et al., 2014), complicating the elucidation of distinct functional roles. Our detection of severe sarcomeric phenotypes following disruption of Fhos activity on its own, identifies Fhos as a key contributor to the processes governing IFM thin-filament array assembly, and sets the stage for studying the involvement of formins in this major aspect of microfilament organization, via utilization of genetic approaches. FHOD-family formins have been previously associated with sarcomere organization in cardiac muscle (Iskratsch et al., 2010; Kan et al., 2012; Taniguchi et al., 2009; Wooten et al., 2013), but the roles these proteins play during skeletal myogenesis have been difficult to ascertain. The ability to dissect the program of IFM sarcomere formation and to disrupt the activity of Fhos to different extents and at distinct phases, provides a comprehensive view of the role of this formin-family protein in the process, and of skeletal muscle thin-filament array assembly at large (Figure 7).

Model of IFM thin-filament array assembly and the roles of Fhos and Sals.

Four distinct stages of thin-filament array assembly and maturation are represented in correlation to pupal developmental stages. (A) Extensive filament polymerization (0–30 hr APF), which takes place in a Fhos-independent manner. (B) Organization of nascent sarcomeres and patched actin incorporation (30–45 hr APF). Fhos localizes to the nascent arrays, and is required for their organization into discrete structural units. (C,D) Growth and maturation of nascent sarcomeres (45 hr APF- eclosion). Fhos localizes to the vicinity of the Z-discs, and is essential for radial growth of the thin-filament arrays, possibly via peripheral filament recruitment. Two additional actin incorporation modes are executed in a Fhos independent manner. Z-disc associated monomer turnover, and 'pointed' end elongation, which is mediated primarily by Sals.

https://doi.org/10.7554/eLife.16540.014

1. The onset of pupal development is accompanied by a prominent burst of actin polymerization, generating a store of microfilaments for use during subsequent stages, when polymerization activity declines considerably. The identity of actin nucleators and in particular formins involved in the extensive initial polymerization is not known, and it is certainly possible that several formins act redundantly, given our failure to disrupt this process by single formin knockdowns, including that of Fhos. An initial sign of internal myofiber organization is seen in the segregation of the abundant microfilaments produced during the early phase of pupal development, into elongated myofibrils that align in parallel to the fiber. We do not know the nature of the signal governing this alignment, but it is noteworthy that previous studies of the dynamic organization of sarcomeres in cultured muscle cells have indicated that microtubules which are aligned along the fiber may provide an initial cue for the recruitment and orientation of myosin heavy chain and possibly microfilaments as well (Pizon et al., 2005).

2. The initial, widespread polymerization gives way to a more limited mode of incorporation, characterized by 'patches' of actin monomers that are added to a nascent microfilament array, suggesting that this period is devoted to proper structuring of the arrays within individual, uniformly-sized and regularly separated sarcomeric units. It is during this interim period that disruption of Fhos activity first leads to alteration of the monomer incorporation pattern, in what we interpret to be a telling fashion, as it retains an underlying, highly regular pattern of monomer incorporation at the pointed-ends of the thin filament arrays (Figure 4D–G). This observation constitutes a direct demonstration that IFM arrays elongate from their pointed-ends, contrary to the conventional barbed end-biased growth of microfilaments, and in keeping with previous findings based on studies of the pointed-end capping protein Tropomodulin in both IFMs and vertebrate cardiomyocytes (Littlefield et al., 2001; Mardahl-Dumesnil and Fowler, 2001). Furthermore, our investigation identifies the WH2-domain protein Sals as a major mediator of the pointed-end growth of IFM thin-filament arrays, similar to its function in Drosophila larval muscle sarcomeres (Bai et al., 2007). Interestingly, Leiomodin acts to mediate pointed-end elongation in vertebrate cardiomyocytes (Chereau et al., 2008; Tsukada et al., 2010), implying a conserved function for WH2-domain actin regulators in sarcomerogenesis.

Interfering with Fhos activity results in severe impairment of myofibril and sarcomere organization (Figure 2), raising the question of how to reconcile the strong mutant phenotypes with the limited degree of Fhos-dependent actin incorporation and array growth during the early and interim periods of pupal development. We suggest that, rather than participating directly in the process of actin polymerization, Fhos plays a key organizational role during this period, which leads to the initial structuring of thin-filament arrays, using microfilaments generated by other formins and actin regulators. FHOD-family formins are considered to be poor microfilament nucleators (Schonichen et al., 2013; Taniguchi et al., 2009), and are thought to act via alternative modes- such as microfilament bundling (Kutscheidt et al., 2014; Schonichen et al., 2013)- to provide shape and structure to microfilament arrays. Our analysis is consistent with this notion, as we have demonstrated that FhosI966A, a Fhos variant presumably lacking barbed end associated activities, supports formation of small but properly organized sarcomeres, similar to the sarcomeres normally formed during the early and interim stages of pupal development. We suggest therefore, that Fhos plays a critical early role, coupling organization of pre-existing filaments into ordered arrays, with a secondary but important capacity to coordinate array size and structure through 'patchy' monomer incorporation.

3. Once the rudimentary sarcomere 'core' is assembled and defined Z-discs with fixed spacing are established, further extension and addition of actin filaments is dictated by the initial organization of the sarcomere. It is at this stage that Fhos assumes a striated localization pattern corresponding to thin-filament array ends, and where localization to the barbed end region becomes critical for Fhos function. We identified three different modes of actin incorporation which contribute to the nascent sarcomere maturation during the later stages of pupal development:

  1. Elongation. Thin-filament arrays continue to extend laterally. Growth occurs from the pointed-ends and is mediated primarily by Sals. On the other hand, no evidence for extension at the barbed ends is evident from actin-GFP incorporation. We suggest that the reduced lateral size of arrays observed following late-stage Fhos knockdown may arise from the compromised protection of the barbed ends immediately adjacent to the Z-disc.

  2. Radial 'thickening'. The sarcomeres grow radially due to the circumferential addition of actin filaments at their periphery. The added filaments are predominantly synthesized at this later stage, since they are readily labeled by monomeric actin produced only at that time (Figure 1E,E’). Radial growth requires Fhos, and therefore constitutes a second major contribution of this formin to thin-filament array assembly. Fhos may contribute to radial thickening by bundling and recruiting complete filaments to the periphery of the array core. The reliance on localization to the vicinity of the Z-discs and the arrest in growth of FhosI966A sarcomeres prior to radial thickening strongly imply that this activity depends on association of Fhos with the barbed ends of the thin-filament arrays.

  3. Barbed-end turnover. An unexpected finding was the identification of rapid actin turnover that is highly restricted to the barbed ends of the thin-filament arrays. The barbed-end turnover persists throughout the late phase of pupariation, and does not contribute to filament elongation (Figure 1G–J). Interestingly, barbed-end turnover could be detected with the GFP-tagged form of the ubiquitous actin isoform actin5C, but not with the IFM-specific isoform GFP-actin88F. This may reflect a structural constraint that underlies the use of distinct actin isoforms for different aspects of array growth and maintenance.

The contribution of barbed-end turnover to thin-filament array organization is not readily apparent. Turnover may be part of a mechanism that maintains the filament integrity by a continuous process of elongation and disassembly within the dynamic environment of the maturing Z-disc. Turnover does not require Fhos, and its molecular basis is currently unknown. It will be interesting to examine if a similar process is operating in adult muscles, to maintain their integrity in the face of extensive contraction activity during flight (Perkins and Tanentzapf, 2014).

In conclusion, our analysis of IFM sarcomeres implies that the assembly of the highly structured thin-filament array is an elaborate, stepwise process involving diverse aspects and machineries of microfilament nucleation, growth and organization. The formin protein Fhos plays a central role, contributing to array assembly at several stages of the process. Fhos acts initially to mediate the assembly of thin-filament arrays within discrete sarcomeric units. Fhos localization to the Z-disc region, an apparently conserved feature among FHOD-family formins (Iskratsch et al., 2010; Mi-Mi et al., 2012; Rosado et al., 2014), becomes essential for function during the later stages of IFM development, where Fhos plays an essential role in sarcomere radial growth. Interestingly, it has been suggested that localization of FHOD3 to the Z-lines of murine cardiomyocytes is a regulated process, relying on phosphorylation of a short domain encoded by an alternatively-spliced exon (Iskratsch and Ehler, 2011; Iskratsch et al., 2010), echoing the importance of Fhos Z-disc localization described here.

While this study has elucidated specific roles for a FHOD-family formin in a model system of skeletal muscle sarcomerogenesis, key issues remain open. The precise molecular nature of the microfilament-associated activities of Fhos, the mechanistic significance of its spatial localization patterns, regulation of its sarcomeric activities and their coordination with other functional elements of the actin-based cytoskeleton, all await further investigation.

Materials and methods

Drosophila genetics

Request a detailed protocol

GAL4 drivers included mef2-GAL4 (Ranganayakulu et al., 1996) and act88F-GAL4 Gajewski and Schulz, 2010], BDSC 38461). UAS-Dicer2 elements were included for enhancement of RNAi activity (Dietzl et al., 2007). GFP-actin lines (Roper et al., 2005) included UAS-GFP-act5C (BDSC # 9258) and UAS-GFP-act88F (BDSC # 9253 and # 9254).

UAS-dsRNA lines used: fhos (VDRC GD2374 [knockdown of long forms] and VDRC KK108388 [general Fhos knockdown]); sals (VDRC KK112869 and TRiP JF01110); tmod (VDRC GD32602 and TRiP JF01094)

Crosses were commonly kept at 25°C. Temporally-controlled expression protocols utilized the GAL80ts/TARGET system (McGuire et al., 2004). F1 progeny were raised at 18°C and shifted to 29°C at 0 hr APF (white pupae). Pupae were then grown at 29°C until the desired developmental time, taking into account the accelerated pupal development (1 hr at 29°C equals approximately 1 hr and 20 min at 25°C) and the time for a complete substitution to inactive form of GAL80 (approximately 5 hr). All indicated time windows are equivalent to the developmental periods for flies grown at 25°C.

Generation of the different fly lines described in the study was achieved as follows:

Fhos-GFP protein trap line: obtained by injection of the protein trap plasmid Splice phase 0 EGFP-FIAsH-StrepII-TEV-3xFlag, into the MiMIC insertion line FhosMI09324, as described (Venken et al., 2011).

UAS-PH-Fhos transgenic line: A full length Fhos-RH cDNA was assembled from clones IP17223 and SD08909 using restriction free cloning as described (Unger et al., 2010). The resulting construct was amplified by PCR, cloned into the NotI–KpnI sites of the pUAST–attB vector containing N-terminal 3XHA and injected following sequence verification into an attP40 line to produce transgenic flies.

Short isoform mutant alleles: A CRISPR/Cas9-based approach (Gratz et al., 2013) was used to target the 1st coding exon of the short Fhos isoforms. A guide RNA template complementary to sequences within the exon (5’CTTCGCCGCCTTCCCGATCCCGGTG3’) was synthesized and cloned into the pU6-BbsI-chiRNA plasmid (Addgene), and the plasmid was injected into vasa-Cas9 embryos (BestGene). Lines were established from the progeny of the injected flies. Mutations were identified by sequencing PCR-amplified genomic DNA encompassing the relevant exon from flies bearing candidate mutant 3rd chromosomes. Two deletion events were identified in this manner, a single guanine nucleotide deletion giving rise to FhosΔG99 (Figure 3A) and a four nucleotide deletion (positions 101–104). Both deletions result in translational frameshifts and premature translational arrest at amino-acid position 58 of the protein sequence, and display identical phenotype and localization features.

Primers used included 5’ GCGTGGCGTGCCAACAATTTG3’ and

5’ GATCGCGATAATGCGATCCACC) for genomic DNA amplification and

5’GTATCTCGTAAATGCGCAG3’ for sequencing.

FhosI966A substitution allele: CRISPR-based genome editing was used to generate the point mutant allele FhosI966A as described (Gratz et al., 2013). Briefly, three 1000 bp fragments, which cover the Fhos FH2 domain genomic region, including exon #20, were synthesized, and cloned into the pHD-DsRed-attB vector (Addgene). The two flanking fragments served as homology arms, while the middle fragment harbored a point mutation leading to substitution of Isoleucine 966 to Alanine. Two different gRNAs 5’CCTCATATAACACCCAATTGGTC and 5’CCATGTAAGAATTAACTTTTGTA) homologous to sites 5’ and 3’ of the replaced genomic area were synthesized and cloned into pU6-BbsI-chiRNA. The plasmid mixture was injected into vasa-Cas9 flies (BDSC#55821) (BestGene). All plasmid constructs were verified by sequencing.

Fhos antibody production

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Antibodies were raised to the 93 C-terminal residues of the Fhos protein. The relevant sequence was amplified by PCR from the SD08909 cDNA clone and cloned into the pDEST17-6xHis vector (Invitrogen). The recombinant protein expressed in BL21 cells, purified and injected into Wistar rats to raise the polyclonal antisera.

Tissue preparation, immunostainings and confocal microscopy

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A modified protocol from (Weitkunat and Schnorrer, 2014) was used for both adult and pupal IFM tissue preparation. Briefly, staged pupae were removed from the pupal case, pinned down on Sylgard plates and dissected in cold relaxation buffer (20 mM phosphate buffer, pH 7.0; 5 mM MgCl2; 5 mM EGTA, 5 mM ATP). For adult IFMs, thoraces of young adults (not older than 48 hr post enclosure) were bisected on the longitudinal axis and collected in the cold relaxation buffer. In both cases fixation was carried out with 4% paraformaldehyde for 20 (pupa) or 30 (adults) minutes at room temp. Following washes and permabilization with PBS+0.3% Triton-X (pupa) and PBS+0.5% Triton-X (adults), the samples were incubated in blocking solution containing 0.1% bovine serum albumin (BSA) + 5% Normal goat serum (NGS). Staining that involved anti-α-actinin and/or anti-MHC required an additional 30 min blocking step with Image-IT FX signal enhancer reagent (Thermo) prior to the standard blocking step. All primary antibodies were diluted in standard blocking solution (0.1% BSA + 5% NGS) and were added for overnight incubation at 4°C. Following washes, secondary antibodies were added for 2 hr at room temp. Adult hemi-thoraces were cleared in 80% glycerol at 4°C overnight prior to mounting. All samples were mounted in Immu-Mount (Thermo).

Primary antibodies and dilutions used included: anti-GFP (chicken, 1:1000, Abcam); anti-MHC (rabbit, 1:1000, kindly provided by P.Fisher, Stony Brook); anti-α-actinin (rat, 1:50, Babraham institute, UK); anti-Zasp52 (rabbit 1:500, [Katzemich et al., 2013]); anti-Obscurin (rabbit, 1:500,[Burkart et al., 2007]); anti-Fhos (rat, 1:200) was generated as described above; anti-Tmod (Rat, 1:500, kindly provided by Velia Fowler).

Secondary antibodies used included Alexa Fluor 405, Alexa Flour 488, Alexa Fluor 555, Alexa Fluor 568 and Alexa Flour 647 conjugated to anti-rabbit, mouse, rat, or chick antibodies (Molecular Probes) and applied at a dilution of 1:1000. Atto647N-Phalloidin (Fluka) was used at 5 μg/ml.

Immunofluorescent images of fixed samples were acquired using Zeiss LSM 710 or Zeiss LSM 780 confocal scanning systems, equipped with a Zeiss Axiovert microscope, and using a ×20 0.8 N.A or ×63 oil immersion 1.4 N.A lenses. The initial image acquisition was performed using the imaging system Zen software.

Transmission electron microscopy

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Thoraces of young adults or isolated IFMs were collected in the ice cold relaxation buffer (20 mM phosphate buffer, pH 7.0; 5 mM MgCl2; 5 mM EGTA, 5 mM ATP). Following 15 min incubation the samples were transferred into 1 mM sodium cacodylate buffer (pH 7.4) containing fixative (4% paraformaldehyde and 2.5% glutaraldehyde). Samples were fixed for 1 hr (IFMs) or 2 hr (adults) at room temp and transferred to 4°C overnight. Samples were washed x3 with sodium cacodylate buffer, post fixed in 1% OSO4 solution for 1 hr at room temp, washed x3 with sodium cacodylate buffer, incubated in 2% aqueous uranyl acetate for 1 hr and washed x3 with distilled water. Samples were taken through an ethanol dehydration series and incubated in propylene oxide (x3, 10 min each). Infiltration was performed with a series of propylene oxide: Epon mixtures, culminating in incubation in 100% Epon (x3, 12 hr each). Infiltrated samples were embedded in plastic moulds (EMS) and polymerized for 48 hr at 60°. Ultra thin sections were cut using diamond knife 35° (Diatome, Switzerland) on a Leica Reichert ultra cut UCT. Sections were post stained with 1% lead citrate and 2% uranyl acetate.

Images were recorded using an FEI T12 spirit BioTWIN transmission electron microscope (TEM) operating at 120KV and equipped with an Eagle 2Kx2K CCD camera (FEI).

Data analysis

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Measurements of various geometric properties of the sarcomere and tagged actin monomers incorporation were performed using Fiji image analysis software. For sarcomere length and width, the Fiji measurement tool was used to draw a vertical line (width) or horizontal line (length) across the Z-disc of a single sarcomere from pointed end to pointed end, using phalloidin staining as a guide. 50 sarcomeres were measured form 7 different flies for each genotype (350 sarcomeres in total). Horizontal lines or polygons were drawn to measure the Z-disc associated incorporation band length (act5C control and following Fhos knockdown) or the incorporation frame area (act88F) respectively. 50 sarcomeres were measured form 4 different flies for each genotype (200 sarcomeres in total). The intensity distribution in Figure 4E and G was measured along 10 μm horizontal profiles starting from a Z-disc and drawn at the middle of the myofibril. The data represent an average of normalized values collected from 15 profiles in 7 different flies (n = 105) for each genotype. Vertical profiles drawn across half sarcomeres was used to measure the GFP intensity distribution in Figure 4J. The data represents an average of normalized values collected from 100 half sarcomeres in 4 different flies (400 sarcomeres in total). To measure the M/Z incorporation intensity ratio in Figure 5 and S5 polygons were drawn around the Z-disc or M-line area. The date represents an intensity measurement from 50 sarcomeres in 5 different flies (250 sarcomeres in total). The F-actin intensity distribution in Figure 5 and S5 was measured along a 13 μm horizontal profile. The data represents an average of normalized values collected from 15 profiles in 5 different flies (n = 75). Distribution of actin incorporation events was obtained from 50 sarcomeres in 5 different flies (250 sarcomeres in total). The incorporation events were visualized by a GFP intensity profile drawn along the contour of the nascent arrays. Z-disc and M-line vicinity markers (Zasp52 and Obscurin, respectively) determined the location of the incorporation event.

For counting thick filaments in TEM cross-sections, a threshold base segmentation was applied and the number of filaments determined by using Fiji Analyze Particles tool. The data represent an average thick filament number per myofibril from 20 myofibrils from 3 different samples.

All graphs and statistic tests were done using GraphPad Prism software. The figures were assembled and organized using Adobe Photoshop CS6.

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Decision letter

  1. Mohan Balasubramanian
    Reviewing Editor; University of Warwick, United Kingdom

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: this article was originally rejected after discussions between the reviewers, but the authors were invited to resubmit after an appeal against the decision.]

Thank you for submitting your work entitled "The Drosophila formin Fhos is a primary mediator of sarcomeric thin-filament array assembly" for consideration by eLife. Your article has been reviewed by three peer reviewers (with expertise in Drosophila cell biology, cytoskeleton, and muscle cell biology), and the evaluation has been overseen by a Reviewing Editor and Vivek Malhotra as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Velia Fowler (Reviewer #2).

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

In discussions, the referees agreed that the description of the stages involved in assembly of thin filaments was very thorough and well done. The referees agreed that the quality of data was very high as well.

However, several questions were raised (please see referee comments verbatim), in particular on the extent of mechanistic advance and the generality of the conclusions, considering previous work on other formins in muscle and the sals.

Reviewer #1:

The organization of actin-based filaments is central to function of sarcomere, the basic functional units of muscle fibres. In Drosophila, indirect flight musculature (IFM) is a well-established skeletal muscle model system for studying assembly and maturation of thin-filament arrays. This study (Arkadi Shwartz et al.) described the role of Fhos, the single Drosophila homolog of the FHOD sub-family of formins, as a primary mediator of indirect flight musculature (IFM) thin filament organization. The authors also identified several phases in the dynamic construction of thin-filament arrays: intensive microfilament synthesis -> assembly of nascent arrays -> elongation primarily from filament pointed-ends -> radial growth of the arrays via recruitment of peripheral filaments -> continuous barbed-end turnover. In addition, the authors showed that the WH2-domain protein Sals that is known to contribute to pointed-end filament elongation, was specifically responsible for pointed-end elongation.

Unfortunately, this manuscript lacks conceptual novelty and is not suitable for general readers of ELife. Other formin proteins such as Diaphanous has already been studied in skeletal muscle model and this manuscript on Fhos, another formin, does not provide significant mechanistic insight into actin-based thin filaments formation in sarcomere. Given that Sals was previously known to contribute to pointed-send filament elongation, the data on Sals in this manuscript also lacks novelty.

Reviewer #2:

This is an interesting study of the role of Drosophila Fhod subfamily formins (Fhos) in thin filament assembly into myofibrils in the Drosophila indirect flight muscle (IFM). The IFM is a great model for sarcomere and thin filament assembly, and the authors take advantage of the fly genetics as well as the ability to express 'pulses' of GFP-actin during IFM myofibril assembly to evaluate the role of the Fhos proteins in actin incorporation/assembly at the barbed vs pointed ends of the thin filaments, as well as assembly of new peripheral thin filaments during stages of myofibril growth. This is an important area in which relatively few careful studies combining genetic deletion or knockdown with tests of actin incorporation have been undertaken. The key findings are that the long isoform of Fhos (Fhos-PH) is required for actin assembly at barbed ends and peripheral thin filament addition during myofibril growth, and that the nucleation activity of Fhos is not essential for this function. In addition, data is presented suggesting that the SALS protein is important for pointed end actin addition and thin filament elongation. The data is of high quality and interesting and reinforce and extend substantially the earlier findings from Mardahl-Dumesnil and Fowler (2001) that thin filament elongation during myofibril assembly takes place at their pointed but not barbed ends, and from Bai et al. (2007) that SALS is important for thin filament elongation at pointed ends. There are a few areas that require clarification and extension to solidify the conclusions.

1) The image of the thin myofibrils in Figure 3D showing partial rescue of the Fhos MI01421 with the GFP-Fhos-PA (short Fhos) is described as having "only weak corrective influence on the disrupted microfilament organization and other sarcomeric defects in myofibrils (Figure 3C,D)". However, to my eye this image shows thin myofibrils but with clearly demarcated and short sarcomeres, quite similar to the image of the myofibrils for the point mutant Fhos I966A in Figure 6A-A', which is described as having "an organized structure of repeated sarcomeric units, with clear demarcation of the Z-discs (Figure 6A-A'), suggesting an arrest in sarcomere growth following proper initial assembly and organization." Thus, it appears to me that the rescue with the short GFP-Fhos-PA should be reinterpreted, and the text rewritten. See next point also.

2) The experiment in Figure 4 shows that the Fhos-i knockdown (which reduces long Fhos isoforms specifically) reduces GFP-actin incorporation at Z lines and at the myofibril periphery, but does not affect the GFP-actin incorporation at the M line. This is a very nice experiment. However, since the short Fhos-PA localizes to the M lines (Figure 3 H-H'), and the Fhos-PA does rescue the myofibril disruption phenotype to some extent (Figure 3D), this could mean that the short Fhos-PA functions at M lines to promote GFP-actin incorporation. However, this was not tested. Depending on the results, the model in Figure 7 may need to be modified.

3) The GFP-actin (act88F) incorporation in the sals RNAi experiment (Figure 5E-E') is not convincingly qualitatively different from the control in Figure 5D-D'. I can see both M and Z line incorporation, just that some M lines appear fainter than others, and overall the image appears to have reduced/fuzzier GFP-actin incorporation. (Note that the shorter sarcomeres due to reduction of sals are convincing (Figure 2—figure supplement 2) and repeat the previous study from Bai et al.) However, I wonder whether the sals is actually affecting pointed end actin elongation directly. Do the authors have stronger data for the GFP-actin incorporation locations? Can the relative M and Z line incorporation be quantified by line scans to compare the WT and the knockdown?

4) Along these same lines, the study of Mardahl-Dumesnil and Fowler implied that Tmod reduces actin pointed end incorporation and thus prevents elongation. However, these authors never tested a Tmod knockdown or deletion, or actin incorporation. Can the authors perform a knockdown of fly Tmod in the IFM to determine whether this affects GFP-actin incorporation at the pointed end? This would fill in the picture, and strengthen the unique role(s) of the Fhos isoforms.

5) In Figure 6, the overall thin/thick filament organization appears normal by TEM of cross sections in the small myofibrils that form in the Fhos I966A point mutant (Figure 6C). However, I noticed a few areas of discontinuity in the lattice (upper left of myofibril) in which there are missing or extra thin filament surrounding the thick filament. In Figure 6B, there is an impression that the M line is not perfectly straight and that H zone is not perfectly even, so that some thin filaments may be shorter and some longer. I wonder whether there may be some subtle thin filament length or organization defects. Can the authors comment on this?

6) In Figure 1, can higher magnification images of the GFP-actin88F for the 45 h incorporation be shown to help identify the location of the actin "patches"? It appears that they do not colocalize with the Z line marker, Zasp. Do they colocalize with the pointed end marker, Tmod? Similarly, in the FhosMI-GFP experiment in Figure 3 F-F', where are the FhosMI-GFP patches? The colocalization with α-actinin is not shown at high magnification. (The regular sarcomeres at later times make it easier to figure out what is going on).

Reviewer #3:

The study describes 3 stages of actin monomer incorporation during Drosophila indirect flight muscle differentiation. Initial bulk polymerisation is followed by ordering into units, which are then elongated and widened to match the uniform sarcomere size. The formin FHOD is implicated in organising and thickening the sarcomere, which requires its long isoforms with N-terminal Z-disk localisation domain. Sals is required for pointed end elongation (as had been shown previously). In both these areas actin88F is incorporated, while at the barbed end, a different actin isoform, actin5c is being built in in a FHOD independent manner.

FHOD seems not to require its barbed-end association (and any associated nucleation, elongation and capping function) until about 60h APF as a conserved mutation I966A can still form organised sarcomeres. Thus the authors argue that FHOD might use its bundling activity during the early stages. However, such a bundling activity is neither shown for FHOD in the present study, nor in the two cited papers.

It is slightly puzzling why FHOD needs to localise to the barbed end, but doesn't need to actually bind to the barbed end. Any mechanistic explanations provided are speculative as no data as to the activity of the mutant or mechanistic action of FHOD or Sals itself are contained in the paper. Thus interpretations rest on the assumption that these would function as other formins. Interestingly, FHOD3, the closest mammalian homologue of FHOD inhibits barbed end elongation, rather than accelerating it (Taniguchi 2009) – a fact that seems not to be discussed in this study.

While the N-terminal extension of the longer FHOD isoforms seem to be of functional importance to organise sarcomeres and the longer isoforms localise specifically to the Z-disk, a causal relationship cannot be drawn, i.e. it is not clear that the localisation itself is required or that the localisation is sufficient to enable the canonical elements of FHOD to fully function. This would need to be tested with more subtle mutations that affect localisation (rather than the removal of 120kD!) and/or using means to bypass the requirement for the N-terminal domain by a more direct recruitment of the shorter FHOD isoforms to Z-disks. In the absence of such mechanistic experiments, the observations are difficult to interpret and some of the statements would need to be altered or stated as being purely speculative.

To my understanding, no data show that radial size increase occurs through recruitment of peripheral filaments – instead incorporation of actin monomers suggest their new synthesis. Also, the barbed end-binding activity of FHOD seems to be required for this step, whether nucleating or inhibitory.

Overall a beautifully illustrated descriptive story identifying temporal and spatial patterns of actin incorporation and formin activity. However, the authors have over-interpreted their findings and make mechanistic claims that are not substantiated by data in the study (or elsewhere).

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for submitting your article "The Drosophila formin Fhos is a primary mediator of sarcomeric thin-filament array assembly" for consideration by eLife.

Your appeal has been reviewed by an additional peer reviewer, and the evaluation has been overseen by a Reviewing Editor and Vivek Malhotra as the Senior Editor. The reviewer has opted to remain anonymous.

While the additional referee sees the value of your description of actin polymerization and thin filament assembly in Drosophila IFM, they have raised a number of points, all of which seem very reasonable and geared towards tightening controls and improving imaging quality.

In summary, we would like you to revise the manuscript, taking into consideration the comments of all the four referees, and in particular on the substantive points raised by referees 2, 3, and 4. You already have the comments of reviewer# 1, 2 and 3, and below are the comments of reviewer #4

Reviewer #4:

Arkadi Shwartz et al. present a study on Drosophila indirect flight muscle actin assembly during the various stages of development and observe the incorporation of different actin isoforms (act88F, act5c) into specific locations of the sarcomere. The authors find that Fhos the single Drosophila homologue of the mammalian FHOD subfamily, which has a documented role in sarcomere assembly and/or maturation, is crucially required for IFM integrity. Knockdown and a previously described mutation lead to thin and randomly oriented myofibrils, with defects appearing after 50hs APF, suggesting an involvement at early stages of sarcomere formation. They further analyze the involvement of different isoforms as well as SALS in the regulation of actin assembly and find that the long isoform is targeted to the Z-disc, where it assembles and bundles the actin, while the short isoform is localized to the pointed ends and dispensable for myofibril formation. SALS, in contrast, as previously demonstrated (Bai et al., 2007) is needed to regulate the thin filament length at the pointed end. The article provides an unprecedented level of detail regarding the different stages of actin assembly during myofibrillogenesis and the involvement of FHOD proteins therein. There are however several points that need to be addressed before publication.

Major points:

1) The data presented regarding the unsuccessful rescue of the Fhos deletion phenotype by FHOS-PA is not entirely convincing. While there is an apparent lack of rescue in Figure 3—figure supplement 1A,A', the stronger overexpression of Fhos-PA results in clearly thicker and more organized sarcomeres in Figure 3D, suggesting that the short isoform can, at least at high expression levels take over the function and even localize to the Z-disc (Figure 3—figure supplement 1C,C'). The data from targeting the 5'coding exon provides further evidence for the involvement of the large isoforms (Figure 3 E,E'), however, the authors should perform rescue experiments with the PH isform as well, to show that it can fully restore the myofibril integrity.

2) Further, regarding the isoforms and the conclusions that the authors draw from their data:

a) The authors suggest that PA and PH are the main isoforms, but no data (qPCR or similar) is presented, or studies cited, to this regards.

b) Disregarding the expression levels, a rescue with the π isoform and localization of the π isoform should be tested as well to exclude major involvement of this isoform.

3) The authors state that the short isoform is inactive, due to its localization to the vicinity of the M-band (3H,H'). This is however not supported by the presented data since the authors find 1) addition of actin to M-Bands, and thickening of myofibrils at late stages of development (Figure 1E); 2) the authors argue, referencing the data from Schoenichen et al. for FHOD1, that Fhos has an important actin bundling activity, which requires filament side binding, however not necessary barbed end binding. 3) stage specific localization changes have been reported for FHOD3 as well, with early localization to the vicinity of the M-Band (Iskratsch et al., 2013). 4) the data from Figure 4D-G demonstrates a patchy incorporation of actin at 60hrs, not only at the Z-disc, which is lost after knockdown of Fhos. This demonstrates that localization to the Z-disc is not necessarily the single determinant of actin assembly activity actin filament turnover could be contributing to differing localization, especially since it was shown in living zebrafish that YFP-actin is incorporated all along the I-Band(Sanger et al., 2009). The authors could test this by performing FRAP experiments to investigate Actin turnover at the Z-disc, around the M-Band and the periphery of the sarcomeres in presence or absence of the various isoforms.

4) Several of the images comparing the control and Fhos deletion/knock downs are difficult to compare due to varying quality, with fuzzier staining also present in the other channels (especially F-actin and zasp in Figure 4 H vs I and J vs K, and F-actin vs Obscurin Figure 5 D vs E). The authors should provide better comparable images (or if not possible, discuss the reason for this difference). Also the authors should provide quantifications of the intensities (e.g. of the M and Z-band intensities normalized to the background).

5) The GFP-act88F localization around the M-bands is narrower in the sals-i, but still present. The authors should provide quantifications for the width (i.e. does this account for the difference in thin filament length) and test or at least discuss what other proteins could be responsible for the ongoing pointed end assembly in absence of sals.

https://doi.org/10.7554/eLife.16540.015

Author response

[Editors’ note: the author responses to the first round of peer review follow.]

Reviewer #1:

The organization of actin-based filaments is central to function of sarcomere, the basic functional units of muscle fibres. In Drosophila, indirect flight musculature (IFM) is a well-established skeletal muscle model system for studying assembly and maturation of thin-filament arrays. This study (Arkadi Shwartz et al.) described the role of Fhos, the single Drosophila homolog of the FHOD sub-family of formins, as a primary mediator of indirect flight musculature (IFM) thin filament organization. The authors also identified several phases in the dynamic construction of thin-filament arrays: intensive microfilament synthesis -> assembly of nascent arrays -> elongation primarily from filament pointed-ends -> radial growth of the arrays via recruitment of peripheral filaments -> continuous barbed-end turnover. In addition, the authors showed that the WH2-domain protein Sals that is known to contribute to pointed-end filament elongation, was specifically responsible for pointed-end elongation.

Unfortunately, this manuscript lacks conceptual novelty and is not suitable for general readers of ELife. Other formin proteins such as Diaphanous has already been studied in skeletal muscle model and this manuscript on Fhos, another formin, does not provide significant mechanistic insight into actin-based thin filaments formation in sarcomere. Given that Sals was previously known to contribute to pointed-send filament elongation, the data on Sals in this manuscript also lacks novelty.

We take issue with the reviewer’s assessment of the novelty of our findings vis-a-vis published studies on Formin function during skeletal muscle sarcomerogenesis. Our study couples a detailed analysis of the dynamics of thin-filament array assembly, at high spatial and temporal resolution, with assignment of distinct functional roles to a single Formin sub-family (FHOD/Fhos). We are unaware of similar comprehensive studies, or of such clear demonstrations of sarcomeric Formin function, and believe that we have provided considerable novel insight, using state-of-the-art tools, to a fundamental and somewhat neglected aspect of skeletal muscle biology. Our analysis of Sals function complements the assignment of functional roles to Fhos, and generalizes the significance of this element to sarcomere formation.

Reviewer #2:

This is an interesting study of the role of Drosophila Fhod subfamily formins (Fhos) in thin filament assembly into myofibrils in the Drosophila indirect flight muscle (IFM). The IFM is a great model for sarcomere and thin filament assembly, and the authors take advantage of the fly genetics as well as the ability to express 'pulses' of GFP-actin during IFM myofibril assembly to evaluate the role of the Fhos proteins in actin incorporation/assembly at the barbed vs pointed ends of the thin filaments, as well as assembly of new peripheral thin filaments during stages of myofibril growth. This is an important area in which relatively few careful studies combining genetic deletion or knockdown with tests of actin incorporation have been undertaken. The key findings are that the long isoform of Fhos (Fhos-PH) is required for actin assembly at barbed ends and peripheral thin filament addition during myofibril growth, and that the nucleation activity of Fhos is not essential for this function. In addition, data is presented suggesting that the SALS protein is important for pointed end actin addition and thin filament elongation. The data is of high quality and interesting and reinforce and extend substantially the earlier findings from Mardahl-Dumesnil and Fowler (2001) that thin filament elongation during myofibril assembly takes place at their pointed but not barbed ends, and from Bai et al. (2007) that SALS is important for thin filament elongation at pointed ends. There are a few areas that require clarification and extension to solidify the conclusions.

1) The image of the thin myofibrils in Figure 3D showing partial rescue of the Fhos MI01421 with the GFP-Fhos-PA (short Fhos) is described as having "only weak corrective influence on the disrupted microfilament organization and other sarcomeric defects in myofibrils (Figure 3C,D)". However, to my eye this image shows thin myofibrils but with clearly demarcated and short sarcomeres, quite similar to the image of the myofibrils for the point mutant Fhos I966A in Figure 6A-A', which is described as having "an organized structure of repeated sarcomeric units, with clear demarcation of the Z-discs (Figure 6A-A'), suggesting an arrest in sarcomere growth following proper initial assembly and organization." Thus, it appears to me that the rescue with the short GFP-Fhos-PA should be reinterpreted, and the text rewritten. See next point also.

These points are well taken, and we have now added new observations and have substantially revised our presentation of rescue and localization data, and our discussion of their significance. A key new experiment is specific disruption of the short Fhos isoforms (via CRISPR-based indels- Figure 3 and Figure 3—figure supplement 1). Using this approach we can now state with confidence that the short isoforms are dispensable and the long isoforms are sufficient for full sarcomeric function. The inability of the short forms to provide significant function when expressed at physiological levels is demonstrated both by the severe phenotypes resulting from specifically disrupting the long isoforms, and by the lack of rescue of null alleles by UAS-Fhos-PA, when driven by armadillo-GAL4. We now show this result in figure panels 3D-D’, as was our original intention. We fully agree with the reviewer that over-expression of the short form results in partial rescue- as we show (Figure 3—figure supplement 1B-B”), such rescue is likely the result of “ectopic” localization of the short form to the Z-disc, and, in fact, corresponds to our view that Z-disc localization is critical for Fhos function.

2) The experiment in Figure 4 shows that the Fhos-i knockdown (which reduces long Fhos isoforms specifically) reduces GFP-actin incorporation at Z lines and at the myofibril periphery, but does not affect the GFP-actin incorporation at the M line. This is a very nice experiment. However, since the short Fhos-PA localizes to the M lines (Figure 3 H-H'), and the Fhos-PA does rescue the myofibril disruption phenotype to some extent (Figure 3D), this could mean that the short Fhos-PA functions at M lines to promote GFP-actin incorporation. However, this was not tested. Depending on the results, the model in Figure 7 may need to be modified.

The Fhos-i knockdown in this figure was based on an siRNA construct targeting ALL isoforms. In any case, we now find (upon specific elimination of the short forms) that the fully functional (endogenous) long isoforms localize to both the Z-disc and M-line regions, while Fhos-PH, an isoform that localizes strictly to the Z-disc and contains only part of the large N-terminal region, provides only partial rescue of null alleles. All this leads us to conclude that Fhos may well function at sites other than the Z-disc (where it is essential), including the M-line. While the nature of these non-Z disc roles is currently unknown, we have modified the text to reflect this notion.

3) The GFP-actin (act88F) incorporation in the sals RNAi experiment (Figure 5E-E') is not convincingly qualitatively different from the control in Figure 5D-D'. I can see both M and Z line incorporation, just that some M lines appear fainter than others, and overall the image appears to have reduced/fuzzier GFP-actin incorporation. (Note that the shorter sarcomeres due to reduction of sals are convincing (Figure 2—figure supplement 2) and repeat the previous study from Bai et al.) However, I wonder whether the sals is actually affecting pointed end actin elongation directly. Do the authors have stronger data for the GFP-actin incorporation locations? Can the relative M and Z line incorporation be quantified by line scans to compare the WT and the knockdown?

We now use fluorescence intensity measurements to provide quantification of both the sarcomere shortening (Figure 5F) and reduced M/Z incorporation ratio (Figure 5G) phenotypes associated with sals knockdown.

4) Along these same lines, the study of Mardahl-Dumesnil and Fowler implied that Tmod reduces actin pointed end incorporation and thus prevents elongation. However, these authors never tested a Tmod knockdown or deletion, or actin incorporation. Can the authors perform a knockdown of fly Tmod in the IFM to determine whether this affects GFP-actin incorporation at the pointed end? This would fill in the picture, and strengthen the unique role(s) of the Fhos isoforms.

Tmod is indeed a logical candidate for an element involved in pointed end elongation. As recommended by the reviewer, we assessed and analyzed the effects of knockdown in similar fashion to the study of sals. In general the tmod knockdown phenotypes are quite mild, and possibly indicate an early role for Tmod in thin-filament array assembly. The relevant observations are shown in Figure S5.

5) In Figure 6, the overall thin/thick filament organization appears normal by TEM of cross sections in the small myofibrils that form in the Fhos I966A point mutant (Figure 6C). However, I noticed a few areas of discontinuity in the lattice (upper left of myofibril) in which there are missing or extra thin filament surrounding the thick filament. In Figure 6B, there is an impression that the M line is not perfectly straight and that H zone is not perfectly even, so that some thin filaments may be shorter and some longer. I wonder whether there may be some subtle thin filament length or organization defects. Can the authors comment on this?

We thank the reviewer for pointing out these imperfections in the Fhos I966A mutant arrays. However, given that these were obtained from 1 day old adult flies, following several days of IFM growth arrest, they may well be the result of secondary effects. We therefore prefer not to speculate about their significance, and to limit our interpretation to the general resemblance they bear to intermediate stage IFMs.

6) In Figure 1, can higher magnification images of the GFP-actin88F for the 45 h incorporation be shown to help identify the location of the actin "patches"? It appears that they do not colocalize with the Z line marker, Zasp. Do they colocalize with the pointed end marker, Tmod? Similarly, in the FhosMI-GFP experiment in Figure 3 F-F', where are the FhosMI-GFP patches? The colocalization with α-actinin is not shown at high magnification. (The regular sarcomeres at later times make it easier to figure out what is going on).

Following these comments, we have extended our analysis of the actin “patches”, by adding magnified panels and fluorescence intensity profiles for 45 hr IFMs stained for GFP-actin, phalloidin and the Z-disc and M-like markers Zasp and Obscurin (Figure 1—figure supplement 1B-E). Further quantification clearly demonstrates that the patches tend to localize to the ends of the arrays (Figure 1—figure supplement 1F).

Reviewer #3:

The study describes 3 stages of actin monomer incorporation during Drosophila indirect flight muscle differentiation. Initial bulk polymerisation is followed by ordering into units, which are then elongated and widened to match the uniform sarcomere size. The formin FHOD is implicated in organising and thickening the sarcomere, which requires its long isoforms with N-terminal Z-disk localisation domain. Sals is required for pointed end elongation (as had been shown previously). In both these areas actin88F is incorporated, while at the barbed end, a different actin isoform, actin5c is being built in in a FHOD independent manner.

FHOD seems not to require its barbed-end association (and any associated nucleation, elongation and capping function) until about 60h APF as a conserved mutation I966A can still form organised sarcomeres. Thus the authors argue that FHOD might use its bundling activity during the early stages. However, such a bundling activity is neither shown for FHOD in the present study, nor in the two cited papers.

We feel that our study covers considerable ground in breaking down the process of thin-filament array assembly into a discrete series of continuous events, some of which overlap temporally, and in assigning functional roles to several key elements that mediate these processes. Admittedly, we do not provide biochemical data that tests the actin-associated activities of Fhos (not for lack of trying- Fhos expression at useful levels has proven to be very difficult). However, we feel compelled to use the ample descriptive and genetic data provided to speculate on molecular mechanisms, bearing in mind established molecular capacities of the proteins involved. That said, we respect the reviewer’s expert assessment that some of the statements come across as over-interpretation of the data, and have tried to reorganize the text and “tone down” the language (in the main text and the relevant mention in the abstract) accordingly.

With regard to FHOD formins acting as microfilament bundling proteins- here we feel fully justified in building upon this notion, which was clearly demonstrated by Schonichen et al. (JCS 2013) using a number of assays, and is treated as an established capacity of FHOD proteins in various papers including the study from Kutscheidt et al. (NCB 2014) which we cite. However, we agree that initial mention of this specific activity was granted excessive prominence, and has now been moved from the Results section to the Discussion. Further mention of a bundling capacity and its significance are now phrased in a less dogmatic fashion.

It is slightly puzzling why FHOD needs to localise to the barbed end, but doesn't need to actually bind to the barbed end. Any mechanistic explanations provided are speculative as no data as to the activity of the mutant or mechanistic action of FHOD or Sals itself are contained in the paper. Thus interpretations rest on the assumption that these would function as other formins. Interestingly, FHOD3, the closest mammalian homologue of FHOD inhibits barbed end elongation, rather than accelerating it (Taniguchi 2009) – a fact that seems not to be discussed in this study.

We fully agree that the critical nature of Fhos localization to the Z-disc region does not readily correspond with its presumed molecular capacities. As brought up in the Discussion, Z-disc region localization appears to be a general feature of FHOD proteins in striated muscle, and resolving this matter is a priority for future studies on sarcomeric FHOD/Fhos function. Clearly, determination of the microfilament-regulating properties of Fhos should be a central aspect of such investigations. Our data only allows us to speculate regarding possible mechanisms, but we feel compelled to do so. Again, we have revised the tone of our discussion to properly reflect its speculative nature. The Taniguchi et al. study is now cited as a reference for the “poor nucleation” capacity of FHOD-family proteins.

While the N-terminal extension of the longer FHOD isoforms seem to be of functional importance to organise sarcomeres and the longer isoforms localise specifically to the Z-disk, a causal relationship cannot be drawn, i.e. it is not clear that the localisation itself is required or that the localisation is sufficient to enable the canonical elements of FHOD to fully function. This would need to be tested with more subtle mutations that affect localisation (rather than the removal of 120kD!) and/or using means to bypass the requirement for the N-terminal domain by a more direct recruitment of the shorter FHOD isoforms to Z-disks. In the absence of such mechanistic experiments, the observations are difficult to interpret and some of the statements would need to be altered or stated as being purely speculative.

We think that we have provided a considerable body of data supporting the notion that Z-disk localization is critical nature:

a) The active long isoforms- which are absolutely necessary for sarcomeric Fhos fuction- localize to this site;

b) The short forms do not localize there under physiological conditions, and completely fail to provide function (although they are fully sufficient for Fhos fuction in all other tissues);

c) Partial rescue by the short form is provided by its ectopic localization to the Z-disc region, under circumstances of over-expression.

We believe that this set of results justifies our conclusions regarding the significance of Z-disc localization. We agree that identification and manipulation of a minimal localization sequence within the long isoform is warranted, and would serve as an important step in refining and understanding the nature of Fhos localization within the sarcomere, but this must be left to future studies.

To my understanding, no data show that radial size increase occurs through recruitment of peripheral filaments – instead incorporation of actin monomers suggest their new synthesis. Also, the barbed end-binding activity of FHOD seems to be required for this step, whether nucleating or inhibitory.

We are in agreement with the reviewer on this point- namely, that newly-synthesized microfilaments constitute a major portion (all?) of the filaments added to the periphery of the sarcomere “core”, and stated as such in the Discussion text. We agree and similarly state that barbed-end binding by Fhos is required for its involvement in radial growth of the array. The question we try to grapple with is why a FHOD-type formin, unlikely to be responsible for synthesis, is still essential for radial growth. Bundling is suggested as a possible mechanism.

Overall a beautifully illustrated descriptive story identifying temporal and spatial patterns of actin incorporation and formin activity. However, the authors have over-interpreted their findings and make mechanistic claims that are not substantiated by data in the study (or elsewhere).

We thank the reviewer both for the positive assessment of our experimental work and for pointing out the shortcomings of our interpretation, which we have tried to revise accordingly.

[Editors’ note: the author responses to the re-review follow.]

The additional referee has kindly provided a detailed critique within a few days. While this referee sees the value of your description of actin polymerization and thin filament assembly in Drosophila IFM, the referee has raised a number of points, all of which seem very reasonable and geared towards tightening controls and improving imaging quality.

Reviewer #4:

Major points:

1) The data presented regarding the unsuccessful rescue of the Fhos deletion phenotype by FHOS-PA is not entirely convincing. While there is an apparent lack of rescue in Figure 3 supplement 1A,A', the stronger overexpression of Fhos-PA results in clearly thicker and more organized sarcomeres in Figure 3D, suggesting that the short isoform can, at least at high expression levels take over the function and even localize to the Z-disc (Figure 3—figure supplement 1C,C'). The data from targeting the 5'coding exon provides further evidence for the involvement of the large isoforms (Figure 3 E,E'), however, the authors should perform rescue experiments with the PH isform as well, to show that it can fully restore the myofibril integrity.

These are all good points, some of which were raised by Reviewer #2 as well (please also see our answers there). With regard to the function of the short isoform- we agree, and now better present the data, which we believe to show that rescue by the short forms is possible, but requires localization to the Z-disc. As for the rescuing capacity of the long forms- the CRISPR-based mutation we have now generated, specifically disrupting the short forms, demonstrates that the long forms are both necessary (as shown before) and sufficient. Rescue by the PH form (Figure 3—figure supplement 1C-C’’’) is partial, but while this construct localizes strictly to the Z-disc region, rescue by the endogenous long isoforms is associated with localization to both the Z-disc and M-line regions (Figure 3—figure supplement 1D). We conclude and now state that while Z-disc localization is critical, M-line localization may be of functional importance as well.

2) Further, regarding the isoforms and the conclusions that the authors draw from their data:

a) The authors suggest that PA and PH are the main isoforms, but no data (qPCR or similar) is presented, or studies cited, to this regards.

The use of the term “main” is in error- it has been changed to “representative”, and we thank the reviewer for pointing it out. Form PA is identical in coding sequence to most other short isoforms, and was used by Bogdan and colleagues in their study of Fhos function, to which we refer throughout. PH was used as a proxy for the long forms, since a corresponding cDNA was available to generate reagents.

b) Disregarding the expression levels, a rescue with the π isoform and localization of the π isoform should be tested as well to exclude major involvement of this isoform.

Our new results now suggest, in fact, that the longer forms (PI and PJ) are the ones to provide full functionality, but unfortunately, we were not successful in attempts to generate matching constructs.

3) The authors state that the short isoform is inactive, due to its localization to the vicinity of the M-band (3H,H'). This is however not supported by the presented data since the authors find 1) addition of actin to M-Bands, and thickening of myofibrils at late stages of development (Figure 1E); 2) the authors argue, referencing the data from Schoenichen et al. for FHOD1, that Fhos has an important actin bundling activity, which requires filament side binding, however not necessary barbed end binding. 3) stage specific localization changes have been reported for FHOD3 as well, with early localization to the vicinity of the M-Band (Iskratsch et al., 2013). 4) the data from Figure 4D-G demonstrates a patchy incorporation of actin at 60hr, not only at the Z-disc, which is lost after knockdown of Fhos. This demonstrates that localization to the Z-disc is not necessarily the single determinant of actin assembly activity actin filament turnover could be contributing to differing localization, especially since it was shown in living zebrafish that YFP-actin is incorporated all along the I-Band(Sanger et al., 2009). The authors could test this by performing FRAP experiments to investigate Actin turnover at the Z-disc, around the M-Band and the periphery of the sarcomeres in presence or absence of the various isoforms.

As noted above, our genetic data now supports the notion that Z-disc/barbed-end associated function is not the entire story. While we do not have solid ideas regarding M-line function of Fhos (pointed-end elongation appears to be Fhos-independent), we stress this additional layer of functional complexity in the text.

4) Several of the images comparing the control and Fhos deletion/knock downs are difficult to compare due to varying quality, with fuzzier staining also present in the other channels (especially F-actin and zasp in Figure 4 H vs I and J vs K, and F-actin vs Obscurin Figure 5 D vs E). The authors should provide better comparable images (or if not possible, discuss the reason for this difference). Also the authors should provide quantifications of the intensities (e.g. of the M and Z-band intensities normalized to the background).

We have now improved data presentation according to the reviewer’s comments:

a) In Figure 4H’, I’ and J we quantified the distribution of actin intensity across the arrays, demonstrating enhanced wildtype incorporation at the periphery, which is gone following Fhos knockdown.

b) In Figure 4M we quantified the width of the incorporation band at the Z-disc, an analysis that demonstrates that this turnover is Fhos independent

In this context we wish to note that- as the reviewer observed- the GFP incorporation signal is generally “fuzzier” reviewer upon fhos knockdown, possibly due to accumulation of excess free GFP-tagged actin monomers in the array vicinity.

5) The GFP-act88F localization around the M-bands is narrower in the sals-i, but still present. The authors should provide quantifications for the width (i.e. does this account for the difference in thin filament length) and test or at least discuss what other proteins could be responsible for the ongoing pointed end assembly in absence of sals.

As requested, we have added quantifications in Figures 5 and Figure 5—figure supplement 1, and have examined the possible contribution of Tropomodulin to pointed end elongation (Figure 5—figure supplement 1). Please also see our answers to Reviewer #2 (points 3 and 4).

https://doi.org/10.7554/eLife.16540.016

Article and author information

Author details

  1. Arkadi Shwartz

    Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    AS, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Nagaraju Dhanyasi

    Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    ND, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  3. Eyal D Schejter

    Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    EDS, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    eyal.schejter@weizmann.ac.il
    Competing interests
    The authors declare that no competing interests exist.
  4. Ben-Zion Shilo

    Department of Molecular Genetics, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    B-ZS, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    benny.shilo@weizmann.ac.il
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4903-8889

Funding

Israel Science Foundation (557/15)

  • Eyal D Schejter
  • Ben-Zion Shilo

Weizmann UK (Joint research program)

  • Eyal D Schejter
  • Ben-Zion Shilo

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

Acknowledgements

We wish to thank our lab manager Shari Carmon and BestGene (Chino Hills, CA) for generating the CRISPR-based short isoform mutants and I966A mutant flies, Gali Housman for help in assessing formin knockdown phenotypes and the WIS Electron Microscopy and Antibody Units for technical support. Reagents were kindly provided by Sven Bogdan (U. Munster), Belinda Bullard (U. of York), Paul Fisher (SUNY Stony Brook), Velia Fowler (Scripps Research Institute), Frieder Schöck (McGill U.), the Babraham Institute (Cambridge), the Drosophila Genomics Resource Center (Indiana U.), the Vienna Drosophila Research Center, the TRiP stock center (Harvard Med. School) and the Bloomington Drosophila Stock Center (Indiana U.). We thank our colleagues in the Shilo lab for their continuous input and encouragement. The authors declare no competing financial interests. This work was supported by grants from the Israel Science Foundation and the Weizmann-UK Joint Research Program to EDS and B-ZS. B-ZS is an incumbent of the Hilda and Cecil Lewis chair in Molecular Genetics.

Reviewing Editor

  1. Mohan Balasubramanian, University of Warwick, United Kingdom

Publication history

  1. Received: March 31, 2016
  2. Accepted: September 15, 2016
  3. Version of Record published: October 12, 2016 (version 1)

Copyright

© 2016, Shwartz et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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