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Muscle-specific stress fibers give rise to sarcomeres in cardiomyocytes

  1. Aidan M Fenix
  2. Abigail C Neininger
  3. Nilay Taneja
  4. Karren Hyde
  5. Mike R Visetsouk
  6. Ryan J Garde
  7. Baohong Liu
  8. Benjamin R Nixon
  9. Annabelle E Manalo
  10. Jason R Becker
  11. Scott W Crawley
  12. David M Bader
  13. Matthew J Tyska
  14. Qi Liu
  15. Jennifer H Gutzman
  16. Dylan T Burnette  Is a corresponding author
  1. Vanderbilt University, United States
  2. University of Wisconsin Milwaukee, United States
  3. Vanderbilt University Medical Center, United States
  4. The University of Toledo, United States
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Cite this article as: eLife 2018;7:e42144 doi: 10.7554/eLife.42144

Abstract

The sarcomere is the contractile unit within cardiomyocytes driving heart muscle contraction. We sought to test the mechanisms regulating actin and myosin filament assembly during sarcomere formation. Therefore, we developed an assay using human cardiomyocytes to monitor sarcomere assembly. We report a population of muscle stress fibers, similar to actin arcs in non-muscle cells, which are essential sarcomere precursors. We show sarcomeric actin filaments arise directly from muscle stress fibers. This requires formins (e.g., FHOD3), non-muscle myosin IIA and non-muscle myosin IIB. Furthermore, we show short cardiac myosin II filaments grow to form ~1.5 μm long filaments that then ‘stitch’ together to form the stack of filaments at the core of the sarcomere (i.e., the A-band). A-band assembly is dependent on the proper organization of actin filaments and, as such, is also dependent on FHOD3 and myosin IIB. We use this experimental paradigm to present evidence for a unifying model of sarcomere assembly.

https://doi.org/10.7554/eLife.42144.001

Introduction

At its core, a sarcomere is composed of ‘thick’ myosin II filaments, and ‘thin’ actin filaments (Figure 1A) (Au, 2004). One sarcomere is measured from Z-line to Z-line, which contain α-actinin 2 (Figure 1A). The proper establishment of cardiac sarcomeres during development and their subsequent maintenance is critical for heart function. Previous studies in cultured myocytes have shown the presence of actin bundles called ‘stress fiber-like structures’ similar in appearance to classic stress fibers (Dlugosz et al., 1984). These stress fibers were often found to be close to the edge of the myocyte with sarcomeres existing further from the edge (Rhee et al., 1994). These studies proposed that the stress fibers served as a template for the formation of sarcomeres (Dlugosz et al., 1984; Rhee et al., 1994; Sanger et al., 2005). The original model that proposed this was called the Templating Model (Dlugosz et al., 1984), and was proposed before it was known these stress fibers contained both non-muscle and sarcomeric proteins (Rhee et al., 1994). Beyond non-muscle myosin IIB (NMIIB), which is present in non-muscle cells, stress fibers in muscle cells contain muscle specific proteins, such as α-actinin, tropomyosin, troponins, and tropomodulin (Almenar-Queralt et al., 1999; Rhee et al., 1994; Sanger et al., 2005). Each of these proteins have non-muscle paralogs, which likely serve similar functions (Bryce et al., 2003; Colpan et al., 2013; Côté, 1983; Gunning et al., 2015; Lim et al., 1986; Sjöblom et al., 2008). Partly in response to the presence of muscle specific proteins in stress fibers, the Templating Model was modified to the ‘Pre-Myofibril Model’ (Rhee et al., 1994; Sanger et al., 2005). Even though these models have different names and are often presented as mutually exclusive, they are very similar in their predictions. Specifically, both models posit an actin bundle that appears structurally similar to a stress fiber will acquire a row of sarcomeres over time to become a ‘myofibril’ (Dlugosz et al., 1984; Rhee et al., 1994; Sanger et al., 2005) (Figure 1A). There is a vast amount of localization data in fixed cardiomyocytes to support these models. However, there is very little dynamic data in live cells that suggests stress fibers give rise to sarcomeres. The strongest dynamic support comes from imaging fluorescently tagged α-actinin 2 in myocytes. Time montages from chick skeletal myotubes showed small puncta of α-actinin 2 adding to pre-existing Z lines (McKenna et al., 1986). Subsequently, a time montage was used to show a similar phenomenon occurring in chick cardiomyocytes (Dabiri et al., 1997).

Figure 1 with 1 supplement see all
Sarcomeres arise directly from Muscle Stress Fiber (MSF) precursors.

(A) Electron microscopy (EM) schematic of a cardiac sarcomere from adult mouse. Electron dense regions on the borders of a sarcomere are Z-discs (Z), while the core of the sarcomere is composed of thin actin filaments and thick myosin II filaments (A). Multiple sarcomeres aligned adjacently form a myofibril (lower mag EM, right). (B) hiCM allowed to spread for 24 hr following plating and imaged with SIM. hiCM has been stained for actin and color coding is a representation of height (Z plane) within the cell following 3D imaging (Z-height, left). Notice the clear stress fiber and sarcomere-like actin organization at the front and rear of the cell in box 1 and 2, respectively. (C) Spread U2OS cell color coded for Z as in Figure 1B, displaying prominent actin arc stress fibers behind leading edge of cell and imaged with SIM. Box 1 shows actin arcs just behind the leading edge of cell, while box 2 shows actin arcs on dorsal surface in cell body (green and blue colored actin), while ventral stress fibers (red colored actin) are on bottom surface of cell. (D) Percentage of hiCMs, U2OS, and HeLa cells with actin arc stress fibers. hiCMs; 1372 cells over three experiments. U2OS; 37 cells over four experiments. HeLa; 186 cells over four experiments. (E) Wide-field time lapse of hiCM transfected with Lifeact-mEmerald to visualize actin. MSF at front of hiCM undergoes retrograde flow and acquires sarcomeres (yellow arrows). (F) Laser-scanning confocal microscopy of hiCM expressing Lifeact-mApple showing MSF to sarcomere transition. hiCM lacks sarcomeres at first time point, and MSF at edge of cell undergoes retrograde flow and acquires sarcomeres (yellow arrows). (G) 3D laser-scanning confocal microscopy of hiCM expressing Lifeact-mApple forming sarcomeres. Note how ventral surface (left montage) contains no sarcomere structures, while sarcomere assembly occurs on the dorsal surface of cell (right montage). Scale Bars; (A) 500 nm high mag (left), 2 µm low mag (right); (B) 10 µm low mag, 5 µm high mag insets; (C) 10 µm low mag, 5 µm high mag insets; (E), (F), (G), 10 µm. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.002

Some in vivo data support the Template/Pre-Myofibril Model, while others do not. In strong support of the Template/Pre-Myofibril Model, static images of chick heart tissue have essentially revealed every structure described in primary cultured chick cardiomyocytes (Du et al., 2008). The presence of NMIIB-containing stress fibers in the cardiomyocytes was particularly clear (Du et al., 2008). NMIIB germline knockout (KO) mice were also reported to have fewer and disorganized sarcomeres via EM (Tullio et al., 1997). On the other hand, several studies have called into question the role of stress fibers in sarcomere assembly. First, several studies examining cardiomyocytes within mouse or chick heart tissue did not find stress fibers containing NMIIB (Ehler et al., 1999; Kan-O et al., 2012; Ma et al., 2009). In addition, a conditional KO mouse that removes NMIIB genetically at P9 apparently still had striated sarcomere structures (Ma et al., 2009). Finally, a conditional heart KO of the other major paralog of NMII, NMIIA, was also reported to have no apparent defects in heart formation (Conti et al., 2004; Conti et al., 2015). Taken together, the lack of clear data showing stress fibers in cardiomyocytes and inconsistencies for a role of NMII in sarcomere assembly calls into question whether the Template/Pre-Myofibril Model is a viable construct for understanding sarcomere assembly (Sanger et al., 2005; Sparrow and Schöck, 2009).

There is further data to suggest that a mechanism other than that described in the Template/Pre-Myofibril model could be driving sarcomere assembly. This alternative model—called the ‘Stitching Model’—is based on the idea that parts of a sarcomere are assembled independently and then brought together (i.e., stitched) (Holtzer et al., 1997; Lu et al., 1992; Sanger et al., 2005). In support of the Stitching Model, studies in Drosophila have shown the presence of small myosin filaments following knockdown (KD) of separate Z-line components (Rui et al., 2010). These data suggest that myosin filaments can assemble independently of Z-lines. Indeed, there are also electron micrographs that appear to show stacks of myosin II filaments (i.e., A-bands) without detectable actin filaments in skeletal muscle (Holtzer et al., 1997; Lu et al., 1992; Sanger et al., 2005). Examination of electron micrographs also supports the idea that bodies containing Z-line components and actin filaments—called ‘I-Z-I’ bodies—could also exist in skeletal muscle without apparent myosin II filaments (Holtzer et al., 1997; Lu et al., 1992; Sanger et al., 2005). Based on this data, it was proposed that stitching could occur through sequential assembly by adding new I-Z-I bodies and myosin II filaments (Holtzer et al., 1997; Lu et al., 1992; Sanger et al., 2005).

The Template/Pre-Myofibril Model and Stitching Model have been proposed to be mutually exclusive explanations of how sarcomeres arise. The Template/Pre-Myofibril Model predicts that multiple sarcomeres will appear approximately simultaneously along the length of a stress fiber, while the Stitching Model would predict that sarcomeres will appear adjacently one by one, sequentially (see original models in (Dlugosz et al., 1984; Holtzer et al., 1997; Rhee et al., 1994)). Here, we leverage our discovery that immature human induced pluripotent stem cell-derived cardiomyocytes (hiCMs) completely disassemble and then reassemble their sarcomeres following plating to test these possibilities. Using this assay, we show that sarcomeres are assembled directly from actin stress fiber templates, and we refer to these stress fibers as Muscle Stress Fibers (MSFs). Our data suggest sarcomere assembly is dependent on the formin actin filament nucleator, FHOD3, non-muscle myosin IIA and non-muscle myosin IIB. Surprisingly, our data do not fully support either the Template/Pre-Myofibril Model or Stitching Model, but rather some aspects of each. As such, we now propose a unified model of sarcomere assembly based on the formation of MSFs and their subsequent transition into sarcomere-containing myofibrils.

Results

Development of an assay to test sarcomere assembly

To address how cardiac sarcomeres are assembled, we used hiCMs as a model system (see Materials and methods) (Takahashi et al., 2007). We first noted the actin filaments in hiCMs, which had spread for 24 hr, had two distinct organizations, muscle stress fibers (MSFs) and sarcomere-containing myofibrils (Figure 1B). Spread hiCMs displayed MSFs at the leading edge and organized sarcomere structures in the cell body (Figure 1B). Strikingly, super-resolution imaging revealed the MSFs in hiCMs resembled a classic actin stress fiber found in non-muscle cells, referred to as actin arcs (Figure 1C and D) (Heath, 1983; Hotulainen and Lappalainen, 2006). Actin arcs are stress fibers on the dorsal (top) surface of the cell that are parallel to the leading edge and stain continuously with fluorescent phalloidin (Figure 1C). Similarly, both MSFs and sarcomeres in hiCMs are on the dorsal surface (Figure 1B). We next sought to test the concept that a MSF obtained sarcomeres as predicted by the Templating/Pre-Myofibril Model.

To test whether MSFs give rise to sarcomeres, we needed to develop a sarcomere assembly assay. We noticed that hiCMs which had been freshly plated (1.5–4 hr post plating) contained no sarcomeres at either the cell edge or cell body, as visualized by SIM (Figure 1—figure supplement 1A). Loss of sarcomere structure was confirmed by visualizing multiple sarcomeric proteins, including actin, beta cardiac myosin II (βCMII), α-actinin 2, and TroponinT (Figure 1—figure supplement 1A). Though hiCMs did not contain sarcomeres at early time points post plating, hiCMs did display MSFs at the cell edge (Figure 1—figure supplement 1A). hiCMs subsequently assembled sarcomere structures over the course of 24 hr (Figure 1—figure supplement 1B). We next sought to test if MSFs template sarcomeres. Indeed, time-lapse microscopy of hiCMs expressing the actin probe Lifeact-mEmerald (Riedl et al., 2008) revealed that MSFs acquire sarcomeres over time, with the first sarcomeres appearing between 4 and 16 hr (Figure 1E and F and Video 1). Not surprisingly, this transition occurs on the dorsal surface of the cell (Figure 1G). Importantly, to visualize sarcomere assembly in live hiCMs, we began our imaging at early time points post plating (i.e., 1.5–4 hr). At these time points, hiCMs do not contain sarcomeres (Figure 1—figure supplement 1A), and this ensured we were visualizing the initial sarcomere assembly event, and not sarcomere rearrangement or reorganization.

Video 1
Actin filaments in a hiCM assembling sarcomeres. hiCM transfected with Lifeact-mApple and imaged with SIM.

MSFs undergo retrograde flow and transition to sarcomere containing myofibrils towards cell body. Lookup table: orange hot. 30.5 by 20.9 µm. Video length: 9.5 hr.

https://doi.org/10.7554/eLife.42144.004

To further characterize our sarcomere assembly assay, we used α-actinin 2, which is a classic marker of Z-lines (Luther, 2009). Endogenous α-actinin 2 localized to both MSFs and sarcomeres (Figure 2A). Small puncta of α-actinin 2 localized to MSFs, while sarcomeres had linear α-actinin 2 which labeled Z-lines (Figure 2A). As has been shown in other systems (Dabiri et al., 1997; Du et al., 2008), the spacing between α-actinin 2 puncta increases during the MSF to sarcomere transition, with the spacing of α-actinin 2 ~ 0.5 µm in MSFs and ~1.7 µm in sarcomeres (Figure 2B). In hiCMs, α-actinin 2 puncta in MSFs alternates with NMII, as has been shown for other systems (Figure 2—figure supplement 1) (Ehler et al., 1999; Hotulainen and Lappalainen, 2006; Rhee et al., 1994; Sanger et al., 2005). Interestingly, the spacing of α-actinin 2 puncta associated with MSFs in hiCMs was very similar to the spacing of α-actinin 4 (i.e., a non-muscle paralog of α-actinin) in actin arcs in U2OS cells (Figure 2C). If MSFs were serving as a template for sarcomeres, we asked whether the α-actinin 2 molecules in MSFs were also being incorporated into the sarcomere structures. Previous data suggest α-actinin 2 puncta join existing Z-lines (Dabiri et al., 1997; McKenna et al., 1986). To test this hypothesis, we utilized a photo-convertible probe, tdEOS, which converts from green to red fluorescence to specifically mark the α-actinin 2 puncta of MSFs (Nienhaus et al., 2006; Wiedenmann et al., 2004). We found that a subset of photo-converted α-actinin 2-tdEOS puncta were indeed incorporated into Z-lines (Figure 2D). Collectively, these results strongly suggest MSFs give rise to sarcomeres in hiCMs.

Figure 2 with 1 supplement see all
α-Actinin 2 spacing and dynamics in hiCMs.

(A) Color coded representation for Z-height of endogenous α-actinin 2 in MSFs (box 1) and sarcomeres (box 2) of hiCM imaged with SIM. Note difference in structure and spacing of α-actinin 2 in MSFs (box 1) and sarcomeres (box 2) (B) Distance between α-actinin 2 structures in MSFs and sarcomeres. MSFs; 14 cells, three experiments, 827 measurements, Sarcomeres; 15 cells, three experiments, 527 measurements. Distance between structures increases as MSFs transition to sarcomeres. (C) Histogram depicting distribution of distances between α-actinin 2 structures in MSFs in hiCMs (top) and α-actinin 4 found in actin arcs of non-muscle cells (bottom). Distribution is similar between cell types. (D) Wide-field montage of photoconversion of α-actinin 2-tdEOS in hiCM. MSFs at leading edge of the cell were photoconverted (green to red) and imaged over time. Montage (middle) depicts α-actinin 2-tdEOS puncta of MSFs (hollow yellow arrow heads) transition into sarcomere structures (middle, right). Scale Bars; (A) 10 µm low mag, 5 µm high mag insets. (B), 10 µM. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.005

Actin retrograde flow in hiCMs and non-muscle cells

We next wanted to further investigate the similarities between MSFs and actin arcs. Actin arc stress fibers in non-muscle cells undergo robust ‘retrograde flow’ away from the edge of the cell as can be seen in U2OS cells, a classic model of mesenchymal migration (Figure 3A and C) (Hotulainen and Lappalainen, 2006; Ponti et al., 2004). Kymography measurements found that actin arcs in U2OS cells moved at ~200 nm/min, in agreement with previously published findings (Figure 3A and C) (Ponti et al., 2004). We found MSFs also underwent retrograde flow (Figure 3B and C). Strikingly, however, kymography revealed MSFs in hiCMs moved significantly slower than actin arcs in U2OS cells (Figure 3C). This was the first indication that actin arcs in non-muscle cells are different than MSFs in hiCMs. We next wanted to define the mechanisms governing MSFs and their acquisition of sarcomeres. As the mechanisms of actin arc formation and maintenance have been well studied (Burnette et al., 2014; Hotulainen and Lappalainen, 2006; Murugesan et al., 2016), we were interested in using our assay to test whether the same mechanisms driving actin arc dynamics were governing MSF dynamics.

Retrograde flow of actin in non-muscle cells and hiCMs.

(A) Still of U2OS cell expressing Lifeact-mEmerald (left) imaged with spinning disk confocal. Kymograph (right) taken from purple line of left image. Note robust movement of actin arc stress fibers (yellow arrow). (B) Still of hiCM expressing Lifeact-mApple (left) imaged with spinning disk confocal. Kymograph (right) taken from purple line of left image. Note slower movement of MSF in hiCM compared to actin arcs in U2OS cell, and stationary nature of sarcomeres (Sar). Gamma image correction of 0.5 was used to display relatively bright (i.e., sarcomeres) and dim (i.e, MSF) structures. (C) Quantification of actin stress fiber translocation rates in U2OS cells and hiCMs. U2OS; 3 cells over three experiments. hiCMs; 12 cells over three experiments. Scale Bars; (A), (B), 10 µM. P-values denoted in graph.

https://doi.org/10.7554/eLife.42144.007

Formins, but not the Arp2/3 complex, are required for MSF-based sarcomere formation

The Arp2/3 complex is well known to be required for actin arc formation in non-muscle cells (Hotulainen and Lappalainen, 2006). To test the role of the Arp2/3 complex during sarcomere assembly, we allowed hiCMs to spread in the presence of CK666, an inhibitor of the Arp2/3 complex (Nolen et al., 2009). Surprisingly, hiCMs allowed to spread in the presence of CK666 formed robust MSFs and sarcomeres comparable to untreated control cells (Figure 4A, inset, and 4B). Cells were quantified as containing sarcomeres if they contained three parallel Z lines in a row each separated from the adjacent Z line by 1 µm – 2.5 µm. An α-actinin 2 localization was defined as a Z line if it was as least 2x the length of the microscope’s resolution limit (see Materials and methods). In addition, the spacing between Z-lines between control and CK666 hiCMs was unchanged (Figure 4C). We also found the retrograde flow of MSFs was unchanged between control and CK666-treated hiCMs (Figure 4D and E). To confirm inhibition of the Arp2/3 complex by CK666, we examined the endogenous localization of the Arp2/3 complex with and without CK666 treatment. The strong localization of the Arp2/3 complex at the edge of control hiCMs was absent in CK666 treated hiCMs (Figure 4F and G). To further confirm this observation, we analyzed the loss of p16b, a subunit of the Arp2/3 complex, from the leading edge of hiCMs via CK666 in live hiCMs. Indeed, hiCMs showed rapid loss of p16b-mEGFP from the leading edge following administration of CK666 (Figure 4H and I). The delocalization of the Arp2/3 complex from the leading edge is consistent with inactivation by CK666, as shown previously in non-muscle cells (Henson et al., 2015). Taken together, our data suggest that the Arp2/3 complex does not need to be localized at the leading edge for sarcomeres to be assembled.

The Arp2/3 complex is not required for sarcomere assembly.

(A) hiCM allowed to spread for 24 hr in the presence of 25 µM CK666, labeled with actin and α-actinin 2 (i.e., Z-lines) and imaged with SIM. Box indicates presence of MSFs. (B) Quantification of percentage of cells with sarcomeres at 24 hr post plating in control and 25 µM CK666. Control: 76 cells, 10 experiments; 25 µM CK666: 41 cells, three experiments. (C) Histogram of distribution of distances between α-actinin 2 Z-lines. Note tight distribution of Z-lines in both conditions. Control: 14 cells, three experiments, 317 measurements. 25 µM CK666: 16 cells, three experiments, 530 measurements. (D) Stills of hiCM expressing Lifeact-mApple pre (top) and post (bottom) addition of 25 µM CK666 and imaged with spinning disk confocal. Kymographs (right) taken from dotted yellow line (left). (E) Rates of retrograde flow of hiCMs depicted as percent change in CK666 from pre-drug condition. 8 cells over three experiments. (F) Localization of the Arp2/3 complex in control (top) and 25 µM CK666 treated (bottom) hiCMs imaged with SIM. Note loss of Arp2/3 at the edge of CK666 treated hiCM. Cells spread for 24 hr in presence of 25 µM CK666 as in Figure 4A (G) Quantification of loss of the Arp2/3 complex from the leading edge of hiCMs. Control; 36 cells over three experiments. 25 uM CK666; 29 cells over three experiments. (H) Live hiCM expressing P16B-mEGFP (a component of the Arp2/3 complex) and imaged with spinning disk confocal. Localization of P16B-mEGFP at leading edge in pre-drug control (top) is acutely lost after addition of 25 µM CK666 (bottom). (I) Quantification of hiCMs displaying localization of the Arp2/3 complex (P16B-mEGFP) pre- and post-25µM CK666 in live hiCMs (as in Figure 4H). 27 cells over three experiments. Scale bars; (A) 10 µm low mag, 5 µm high mag inset. (D), (F), (H), 10 µm. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.008

In addition to the Arp2/3 complex, formin-mediated actin polymerization has been shown to be crucial for actin arc formation and dynamics in multiple cell types (Hotulainen and Lappalainen, 2006; Murugesan et al., 2016). As a starting point to test whether formins are required for sarcomere assembly, we allowed hiCMs to spread in the presence of a pan-inhibitor of formin-mediated actin polymerization, small molecule inhibitor of formin homology domain 2, SMIFH2 (Rizvi et al., 2009). SMIFH2 has been shown to stop formin-mediated actin polymerization, actin arc formation and retrograde flow in non-muscle cells (Henson et al., 2015; Murugesan et al., 2016; Rizvi et al., 2009). We found hiCMs spreading in the presence of SMIFH2 completely failed to form sarcomeres (Figure 5A and B, and Figure 5—figure supplement 1). This effect was reversible, as sarcomeres formed after the removal of SMIFH2 (Figure 5—figure supplement 2). Distances between α-actinin 2 structures were also significantly decreased in hiCMs treated with SMIFH2, with the distribution of α-actinin 2 more closely resembling MSFs than sarcomeres (Figures 5C and 2C). However, the alignments of the α-actinin 2 puncta were not similar to MSFs in control hiCMs, as they were not periodic (Figures 5A and 2A). This result strongly suggested formins are required for sarcomere assembly.

Figure 5 with 2 supplements see all
Formins are required for sarcomere assembly and MSF dynamics.

(A) hiCM allowed to spread in the presence of 25 µM SMIFH2 for 24 hr, labeled with actin and α-actinin and imaged with SIM. Box indicates loss of transverse MSFs behind leading edge of hiCM. (B) Quantification of percentage of cells with sarcomeres at 24 hr post plating. Control: 76 cells, 10 experiments; 25 µM SMIFH2, 16 cells, three experiments (C) Histogram of distribution of distances between α-actinin 2 Z-lines. Control: 14 cells, three experiments, 317 measurements, 25 µM SMIFH2: 11 cells, three experiments, 468 measurements. (D) Stills from live hiCM expressing Lifeact-mApple which were spread for 24 hr and assembled sarcomeres. hiCM before (left) and 90 min following addition of 25 µM SMIFH2 (right) (drug administered 24 hr after spreading) and imaged with spinning disk microscopy. Note how sarcomeres and overall actin architecture remains unperturbed at 90 min post 25 µM SMIFH2. (E) Kymographs of MSF and sarcomere retrograde flow taken from purple line in (D). Note immediate loss of retrograde flow following addition of 25 µM SMIFH2. (F) Quantification of actin retrograde flow in hiCMs pre and post addition of 25 µM SMIFH2. Control: 12 cells, three measurements from each cell, three experiments; 25 µM SMIFH2, 12 cells, three measurements from each cell, three experiments. Scale Bars: (A) 10 µm low mag, 5 µm high mag inset. (D), 10 µM. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.009

We next asked if formin inhibition was affecting either the MSFs or sarcomeres directly. To test this, we allowed hiCMs to spread for 24 hr (after they have established sarcomeres) and imaged their actin cytoskeleton via live-cell microscopy before and after administering SMIFH2 (Figure 5D). Following addition of SMIFH2, formation of new MSFs was immediately blocked, along with retrograde flow of existing MSFs (Figure 5D–5F). However, we did not detect any changes in sarcomere structure over the short time of the experiment, and hiCMs continued to beat in the presence of SMIFH2 (note sarcomere structure in Figure 5D). As there are 15 mammalian formin genes, we next asked what specific formin was required for sarcomere assembly.

We performed RNA sequencing analysis of mRNA isolated from hiCMs. Normalized read counts revealed that one formin, FHOD3, was expressed higher than all other formins (Figure 6A). Indeed, previous data from isolated rat cardiomyocytes have shown FHOD3 as crucial for sarcomere maintenance (Iskratsch et al., 2010; Kan-O et al., 2012; Taniguchi et al., 2009). Rat cardiomyocytes containing myofibrils subsequently lost their myofibrils following FHOD3 knockdown (Iskratsch et al., 2010; Taniguchi et al., 2009). However, the role of FHOD3 during de novo sarcomere assembly has not been tested. Therefore, we sought to use our assay to directly test if the formin FHOD3 was required for MSF based sarcomere assembly. We knocked down FHOD3 using siRNA, and hiCMs were unable to assemble sarcomeres following plating (Figure 6B and C). Interestingly, KD of the two most highly expressed formins after FHOD3, DAAM1 and DIAPH1, did not stop sarcomere assembly (Figure 6C and Figure 6—figure supplement 1). However, there are clear defects in the actin organization at the cell edge and in the sarcomeres (Figure 6—figure supplement 1). As FHOD3 had the most prominent phenotype, we decided to focus our further analysis on this condition. In line with pan-formin inhibition, the actin organization and spacing between α-actinin 2 in FHOD3 KD hiCMs highly resembled hiCMs spread in the presence of SMIFH2 (Figure 5A–5C and Figure 6B–6D).

Figure 6 with 1 supplement see all
Formin FHOD3 is required for sarcomere assembly.

(A) Normalized Frames Per Kilobase Million (FPKM) of mRNA expression of the top three expressed formins in hiCMs, three experiments, three separate runs. (B) Actin of siRNA scramble control and siRNA FHOD3 hiCMs allowed to spread for 24 hr and imaged with SIM. Note loss of sarcomeres comparable to SMIFH2 treatment in FHOD3 KD hiCMs (Figure 5A). Western blot (right) denotes protein loss in FHOD3 KD. (C) Quantification of percentage of cells with sarcomeres at 24 hr post plating in scramble control, siFHOD3, siDAAM1, and siDIAHP1 hiCMs. Control: 76 cells, 10 experiments; siRNA FHOD3: 33 cells, three experiments; siRNA DAAM1: 29 cells, three experiments; siRNA Dia1: 26 cells, three experiments. (D) Histogram of distribution of distances between α-actinin 2 Z-lines. Control: 14 cells, three experiments, 317 measurements; siFHOD3: 15 cells, three experiments, 488 measurements. (E) hiCM transfected with FHOD3-mEGFP, fixed at 24 hr, stained for Actin and imaged with SIM. Boxes 1-6 depict localization of FHOD3-mEGFP along MSFs (boxes 1 and 2) and in sarcomeres (boxes 4, 5, and 6). (F) Boxes from (E) showing increased organization of FHOD3-mEGFP from MSFs to sarcomeres. Note increasingly organized structure, and localization between Z-lines of FHOD3-mEGFP. Scale bars; (B), (E), 10 µm. (F), 5 µm. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.012

Based on our results, if FHOD3 is involved in sarcomere assembly, it should localize to MSFs. FHOD3-mEGFP localized to both MSFs at the edge of hiCMs, and then becomes increasingly organized away from the leading edge of the cell where sarcomeres are located (Figure 6E). This localization is consistent with a role for FHOD3 in mediating the transition from MSFs to sarcomeres. Taken together, our data show that the formin FHOD3 localizes to both MSFs and sarcomeres, and is required for de novo sarcomere assembly. We next wanted to investigate other potential mechanisms regulating sarcomere assembly.

Non-muscle myosin II is required for cardiac sarcomere actin filaments

In addition to actin nucleators, non-muscle myosin II (NMII) activity has been shown to be required for actin arc formation and organization in non-muscle cell types (Hotulainen and Lappalainen, 2006; Medeiros et al., 2006). Thus, we asked whether NMII was required for MSF formation and/or the MSF to sarcomere transition. We first localized the two major paralogs of NMII in humans, NMIIA and NMIIB, in spread hiCMs (Vicente-Manzanares et al., 2009). Both NMIIA and NMIIB localize to actin arcs in non-muscle cells (Kolega, 1998). Consistent with this, both NMIIA and NMIIB localized to MSFs, and were restricted from the middle of the cell where sarcomeres were localized (Figure 7A and B). Indeed, time-lapse microscopy revealed NMIIA filaments formed at the edge of hiCMs and underwent retrograde flow as in non-muscle cells (Figure 7C). However, NMIIA remained at the edge of hiCMs and was restricted from the cell body where sarcomeres are formed (Figure 7C and Video 2). NMIIB also remained at the edge of hiCMs and was restricted from the cell body where sarcomeres are formed (Video 3). The vast majority of NMIIA and NMIIB filaments overlapped, except at the very leading edge where NMIIA is localized slightly ahead of NMIIB in hiCMs (Figure 7A and B). Super-resolution microscopy revealed that most NMII filaments contained NMIIA and NMIIB (Figure 7D). NMIIA and NMIIB co-filaments have previously been reported in non-muscle cells (Beach et al., 2014; Shutova et al., 2014). Measurements of the lengths of NMII co-filaments in hiCMs showed lengths agreeing with previously published measurements in non-muscle cells (Figure 7D and E) (Beach et al., 2014; Shutova et al., 2014). Taken together, these data suggest NMII organization and dynamics appear similar in MSFs of hiCMs as in actin arcs of non-muscle cells.

NMII Localization and Dynamics in hiCMs.

(A) Localization of endogenous NMIIA (left) and NMIIB (right) in the same hiCM and imaged with SIM. Both NMIIA and NMIIB localize to MSFs at the leading edge of hiCMs. (B) Line scans starting from edge of hiCMs showing localization of NMIIA (black) and NMIIB (red). Note NMIIA is localized slightly,~1 µm in front of NMIIB. NMIIA: 15 cells, two experiments; NMIIB: 32 cells, four experiments. (C) Color projection of time-lapse of hiCM expressing NMIIA-mEmerald and imaged with laser-scanning confocal. Note how NMIIA-mEmerald remains at the edge of hiCMs. (D) hiCM transfected with NMIIA-mEmerald (N-terminal motors), stained for endogenous NMIIB C-terminal rod domain (cartoon schematic and middle left), and imaged with SIM. High-mag views of NMIIA-NMIIB co-filaments (right) from yellow box (middle left). High mag view of single NMIIA-NMIIB co-filament (bottom) and line scan across white dotted line, from N-terminal motors (purple) and C-terminal rod domains (green). (E) Quantification of NMII co-filament length. Histogram displays the distribution of NMII co-filament lengths (motor-domain to motor-domain). Scale Bars; (A), (C), 10 µm, (D) 10 µm low mag (left), 2 µm ‘zoomed view’ inset (right), 200 nm ‘high mag filament’ inset (bottom).

https://doi.org/10.7554/eLife.42144.014
Video 2
NMIIA filament dynamics during sarcomere assembly. hiCM transfected with NMIIA-mEmerald and Lifeact-mApple and imaged with 3D laser-scanning confocal.

Note how NMIIA-mEmerald filaments form at the edge of hiCMs, are localized to MSFs, but are restricted from sarcomeres during sarcomere assembly. 28 by 35 μm. Video length: 14 hr.

https://doi.org/10.7554/eLife.42144.015
Video 3
NMIIB filament dynamics during sarcomere assembly. hiCM transfected with pHalo-NMIIB and α-actinin 2-mEmerald and imaged with wide-field microscopy.

Note how NMIIB-Halo filaments form at the edge of hiCMs, are localized to MSFs, but are restricted from sarcomeres during sarcomere assembly. 47 by 27 μm. Video Length: 7 hr.

https://doi.org/10.7554/eLife.42144.016

Given the presence of both NMIIA and NMIIB in each filament within MSFs, we next asked whether NMIIA and/or NMIIB were required for sarcomere assembly. NMIIA has previously been shown to be required for actin arc assembly in non-muscle cells (Figure 8—figure supplement 1) (Burnette et al., 2014; Fenix et al., 2016). Thus, we hypothesized that NMIIA would likely be the key paralog required for MSF formation and subsequent sarcomere assembly. Surprisingly, KD of NMIIA did not result in a complete inhibition of MSF or sarcomere assembly, although the sarcomeres in NMIIA KD hiCMs were disorganized (Figure 8A–8E and Figure 8—figure supplements 2 and 3). Notably, NMIIA KD hiCMs displayed a similar distribution of distances between α-actinin 2 structures compared to control hiCMs (Figure 8D). This measurement shows that though there are fewer and more disorganized sarcomeres in the NMIIA KD, their widths as measured from Z-line to Z-line are similar to control hiCMs. However, NMIIA KD cells had significantly shorter Z-lines compared to control hiCMs (Figure 8E). Taken together, this data suggests NMIIA is involved in sarcomere assembly and organization.

Figure 8 with 6 supplements see all
NMIIA and NMIIB are Required for Sarcomere Assembly in hiCMs.

(A) Actin of representative scramble control (top), NMIIA KD (siRNA MYH9, middle), and NMIIB KD (siRNA MYH10, bottom) hiCMs allowed to spread for 24 hr and imaged with SIM. NMIIA KD hiCMs (middle) display disorganized sarcomeres, while NMIIB KD hiCMs (bottom) display no actin-based sarcomeres. (B) Representative western blots of 2 separate experiments showing knockdown of NMIIA (siRNA MYH9, top) and NMIIB (siRNA MYH10, bottom). (C) Percentage of scramble control, NMIIA KD (siRNA MYH9), and NMIIB KD (siRNA MYH10) hiCMs with actin based sarcomeres at 24 hr spread. Control: 49 cells, six experiments; NMIIA KD: 34 cells, three experiments; NMIIB KD: 59 cells, four experiments. (D) Histogram of distribution of α-actinin 2 structures in scramble control, NMIIA KD (siMYH9) and NMIIB KD (siMYH10) hiCMs. siScrambled: 554 measurements, 14 cells, three experiments; siMYH9: 332 measurements, 15 cells, three experiments; siMYH10: 772 measurements, 15 cells, three experiments. (E) Quantification of Z-line lengths in scramble control and NMIIA KD (siMYH9) hiCMs. Control: 22 cells, four experiments; NMIIA: 14 cells, three experiments. (F) Quantification of hiCMs with sarcomeres in scramble control, NMIIA KD (siMYH9), and NMIIB KD (siMYH10) hiCMs before re-plating. siScrambled: 772 cells, four experiments; siMYH9: 642 cells, two experiments; siMYH10: 385 cells, two experiments. Scale bar; (A) 10 µm. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.017

It has previously been shown NMIIB is not required for actin arc formation in non-muscle cells (Kuragano et al., 2018; Shutova et al., 2017) (Figure 8—figure supplement 1). We hypothesized that NMIIB would not be required for MSFs or sarcomere assembly in hiCMs. Surprisingly, NMIIB KD resulted in a complete inability of hiCMs to form sarcomeres after plating (Figure 8A–8D and Figure 8—figure supplements 2 and 4). In addition, the width between α-actinin 2 structures was significantly smaller than control hiCMs (Figure 8D). These results argue NMIIB is also a major player required for sarcomere assembly in hiCMs. To further confirm that myosin II is required for sarcomere assembly, we then pharmacologically inhibited all myosin II paralogs in hiCMs with blebbistatin (Straight et al., 2003). hiCMs spreading in the presence of blebbistatin were unable to assemble sarcomere structures (Figure 8—figure supplement 5). While these defects in sarcomere assembly were dramatic, we noticed that hiCMs treated with siRNA against NMIIA or NMIIB were still beating before re-plating. This implied that the pre-existing sarcomeres of the hiCMs were still intact after KD before plating. Therefore, we immuno-localized α-actinin 2 to visualize sarcomeres in hiCMs before plating. Surprisingly, we found there were no differences between control, NMIIA, and NMIIB KD cells before plating (Figure 8F and Figure 8—figure supplement 6) Collectively, this data would suggest NMIIA and NMIIB are required for de novo sarcomere formation, but not homeostasis (i.e., turnover) of pre-existing sarcomeres.

NMIIB and FHOD3 are required for organized A-band formation

Thus far, our results highlight the importance of formin-mediated actin polymerization and NMII for proper actin filament architecture during sarcomere assembly. We next wanted to address how the thick, β Cardiac Myosin II (βCMII) filaments at the core of the sarcomere (i.e., A-band, Figure 1A) assemble. Therefore, we started by localizing endogenous βCMII and NMIIB filaments (Figure 9A). βCMII predominately localized behind NMIIB in organized sarcomere structures and showed a peak localization ~15 microns behind the leading edge of the cell, with a slight area of overlap with NMIIB (Figure 9A and B). We noted that the area of overlap contained NMIIB-βCMII co-filaments (Figure 9C and D, and Figure 9—figure supplement 1). In addition, we also found NMIIA-βCMII co-filaments in hiCMs (Figure 9—figure supplement 2). To our knowledge, this is the first time a myosin II filament-species has been reported that contains a non-muscle and muscle paralog inside cells. Furthermore, we also found NMIIB-βCMII co-filaments in mouse and human heart tissue, indicating NMIIB-βCMII co-filaments are present in vivo (Figure 9C and Figure 9—figure supplement 1). The co-filaments containing NMIIB and βCMII were of similar length to NMIIA/B filaments (Figure 9D). Indeed, we noticed that near the leading edge of the cell, βCMII filaments are typically smaller and not organized into stacks resembling A-bands (Figure 9A and C–E). This suggests βCMII filaments are polymerized at the edge and subsequently grow larger as they move away from the leading edge (Figure 9E). The presence of NMII before βCMII filaments grow into larger filaments led us to test the hypothesis that NMII would play a role in βCMII filament formation.

Figure 9 with 4 supplements see all
β Cardiac Myosin II (βCMII) Filament Assembly in hiCMs.

(A) Endogenous localization of NMIIB (left) and βCMII in the same hiCM and imaged with SIM. (B) Averaged line-scans of NMIIB (Figure 7B) and βCMII localization in hiCMs spread for 24 hr. Note the peak fluorescence of βCMII is more towards the cell body than peak fluorescence of NMIIB. 23 cells from four experiments were used for βCMII localization. (C) Schematic (top) of NMIIB-βCMII co-filaments. High-mag views of βCMII-NMIIB co-filaments (bottom). Endogenous staining of hiCM (bottom, left) for βCMII (N-terminal motors) and NMIIB (rod domain) and imaged with SIM. Mouse and human tissue (bottom middle, and bottom right, respectively) stained for βCMII (motors) and NMIIB (rod domain) and imaged with SIM and Zeiss 880 with Airyscan, respectively. (D) Histograms displaying width of NMII filaments (top), βCMII filaments (middle), and NMIIB-βCMII co-filaments in hiCMs. Measurements made from motor-domain to motor domain as in Figure 7D and E. (E) Histograms displaying distribution of βCMII filaments widths with respect to their location in hiCMs. Note βCMII filaments tend to grow larger as they move towards the center of the cell. Measurements were not taken from ‘mature’ sarcomere structures in highly organized A-bands. (F) Actin and βCMII of scramble control hiCM (top) and NMIIB KD (siMYH10) hiCM (bottom) spread for 24 hr. Note loss of organized A-bands but presence of βCMII filaments in NMIIB KD hiCM. (G) Fourier transforms of βCMII signal from white boxes in Figure 11F from scramble control and NMIIB KD hiCMs (above and below respectively). Yellow arrows indicate sarcomeric periodicity in scramble control hiCMs, which is lacking in NMIIB KD cells. (H) Actin and α-actinin 2 localized in a hiCM after NMIIB KD. (I) High mag views of actin and βCMII in NMIIB KD (siRNA MYH10) hiCM imaged with SIM. βCMII filaments localize to residual actin filaments. (J) βCMII in hiCM spread for total of 24 hr, with the final 6 hr in 5 µM Latrunculin B and imaged with SIM. Notice lack of βCMII A-bands in the periphery of hiCM, and large βCMII filament aggregates (yellow box). (K) Percentage of scramble control, NMIIB KD (siRNA MYH10), and 5 µM Latrunculin B hiCMs with βCMII A-bands. Control: 26 cells, three experiments; NMIIB KD: 26 cells, two experiments; Latrunculin B: 11 cells, three experiments. Cell bodies were not analyzed in the latrunculin experiment due to the density of βCMII localization. Scale Bars; (A) 10 µm, (C) 200 nm, (F), (G) 5 µm, (H) 1 µm. (I) 10 µm low mag, 5 µm high mag insets. (J) 10 µm low mag, 5 µm high mag inset. P-values denoted in graphs.

https://doi.org/10.7554/eLife.42144.024

To test if NMIIB was also required for βCMII filament and A-band formation, we depleted hiCMs of NMIIB and localized βCMII 24 hr after plating. Compared to control hiCMs, NMIIB KD-hiCMs displayed a significant decrease in the ability to form A-band-like structures (as defined in the Materials and methods) and a reduced overall number of βCMII filaments (Figures 9F, G, I and K). Although βCMII filaments formed, they were highly disorganized compared to control cells, as assessed by Fourier transform (Pasqualini et al., 2015) (Figure 9G). As the actin puncta left over after NMIIB KD contained α-actinin 2 (Figure 9H), we hypothesized that these puncta could also be bound to βCMII filaments. Indeed, we found that βCMII filaments spanned the distance between closely spaced puncta (Figure 9I). This data suggested that even when the actin cytoskeleton is severely disrupted, there are still mechanisms leading to the association between it and βCMII filaments. We next sought to expand upon this observation and test whether the observed defects in βCMII filament assembly in the NMIIB KD hiCMs were caused by the disruption of the actin cytoskeleton. To test this, we allowed hiCMs to form sarcomeres for 18 hr, then treated hiCMs with the actin monomer sequestration agent Latrunculin B for 6 hr (Spector et al., 1983; Wakatsuki et al., 2001). Previous studies in non-muscle cells have shown that latrunculin treatment clears the actin from most of the cell and leaves aggregates of actin filaments scattered throughout the cytoplasm (Ayscough et al., 1997; Gronewold et al., 1999). We also found aggregates of actin filaments in the periphery of hiCM treated with latrunculin (Figure 9—figure supplement 3). βCMII filaments appeared to exclusively localize to these aggregates but were not in organized into A-bands (Figure 11I J). Taken together, these results argue that the organization of actin filaments is a major factor in the organization of βCMII filaments. As FHOD3 KD also resulted in severely disorganized actin filament architecture, we localized βCMII in this condition. Indeed, FHOD3 KD hiCM also had disorganized βCMII filaments compared to control hiCMs (Figure 9—figure supplement 4).

βCMII filaments concatenate to form larger A-band structures.

(A) Cartoon of βCMII filament (left, above). N-terminal tagged human βCMII-mEGFP filament expressed in hiCM (left, below). Gap in signal represents bare-zone lacking motors (green arrows). βCMII single filament and A-band filament (βCMII filaments found within organized A-bands) widths measured by line scans (right). βCMII Filaments: 16 filaments, three experiments. βCMII myofibrils: 28 myofibrils, three experiments. Note more level ‘plateau’ of signal from motors in A-band βCMII filaments. (B) SIM of representative βCMII-mEGFP myofibril in hiCM (top) and laser scanning confocal (bottom). (C) Representative montage showing two separate concatenation events. Yellow arrowhead denotes a large stack of βCMII-mEGFP filaments concatenating with a smaller stack of βCMII-mEGFP filaments as they undergo retrograde flow. Green arrowhead denotes smaller βCMII-mEGFP filament concatenating with larger βCMII-mEGFP stack as they undergo retrograde flow. Both events result in larger and more organized βCMII-mEGFP filament stack (i.e. the A-band). (D) Example of βCMII-mEGFP filament splitting event. Note how small βCMII-mEGFP stack splits to create two smaller βCMII-mEGFP filaments and does not result in larger or more organized βCMII-mEGFP filament stacks. (E) Quantification of % of hiCMs which display concatenation or expansion events of βCMII filaments. Scale Bars; (A) 200 nm, (B), (C) and (D) 2 µm. P-values denoted in graph.

https://doi.org/10.7554/eLife.42144.029

To further investigate the mechanisms of A-band assembly, we created a full length, human βCMII construct containing a mEGFP tag on the motor domain (i.e., N-terminal) (Figure 11A). This construct properly integrated into both single filaments and more mature myofibrils (Figure 11A B). In non-muscle U2OS cells, A-band-like stacks of NMIIA filaments are often formed through a process called ‘Expansion’ (Fenix et al., 2016). During Expansion, NMIIA filaments that are close to each other (i.e., in a tight bundle) move away from each other in space but remain part of the same ensemble, where they can be aligned in a stack similar to muscle myosin II in the A-band. In addition to Expansion, NMIIA filaments also, but more rarely, ‘Concatenated’ (Fenix et al., 2016). Concatenation is defined by spatially separated NMIIA filaments moving towards one another to create a stack. To test how βCMII filament stacks form, we repeated our live-cell sarcomere formation assay using our βCMII-mEGFP construct. In contrast to our previous results with NMIIA, we found the major physical mechanism of βCMII filament stack formation to be concatenation, where pre-existing βCMII filaments ran into one another and stitched together to form the A-band (Figure 11C E and Video 4). A small percentage of hiCMs showed an expansion event of βCMII-mEGFP, however this was significantly less frequent than in non-muscle cells and did not appear to result in a more organized A-band (Figure 11D E). Indeed, each of the hiCMs quantified in Figure 11E showed only one expansion event.

Video 4
βCMII filaments in a hiCM assembling sarcomeres. hiCM transfected with βCMII-mEGFP and imaged with SIM.

Note filaments concatenating. Lookup table: orange hot. 30.7 by 29.7 μm. Video length: 7.5 hr.

https://doi.org/10.7554/eLife.42144.030

Discussion

The goal of this study was to address a decades-old question concerning the origin of sarcomeres assembling in cardiomyocytes. A number of labs have proposed sarcomeres arise from a stress fiber-like precursor, while others propose models where separate sarcomere components sequentially stitch together to form sarcomeres (Holtzer et al., 1997; Lin et al., 1994; Lu et al., 1992; Rhee et al., 1994; Rui et al., 2010; Sanger et al., 2005). Based on previous actin filament staining and NMII localization to these stress fibers, our initial hypothesis was that the mechanisms underlying actin filament polymerization and organization would be related to those governing the non-muscle stress fibers known as actin arcs (see model in Figure 12). In support of this hypothesis, both previous reports in non-human cardiomyocytes and our own work with hiCMs show that the actin organization of these precursors appears identical to actin arcs (Heath, 1983; Rhee et al., 1994; Sanger et al., 2005; Tojkander et al., 2012). Indeed, both MSFs and actin arcs contain NMII, stain continuously with phalloidin, are on the dorsal surface of the cell, and display retrograde flow in which they move away from the cell’s edge (Figure 12). However, our study also revealed distinct differences between the regulation and dynamics of MSFs and actin arcs.

Figure 12 with 1 supplement see all
Model of actomyosin stress fiber formation in non-muscle cells and human cardiomyocytes.

(A) Actin and myosin II stress fiber formation in non-muscle cells. Actin stress fibers are formed via the Arp2/3 complex and the formin mDia1. NMIIA is the predominant isoform at the leading edge of non-muscle cells, and stress fiber formation is NMIIA dependent. Non-muscle cells display robust retrograde flow of actin stress fibers and display rapid turnover. Large NMIIA stacks are formed via growth and expansion of smaller NMIIA filaments. Citations leading to this model are presented in the cartoon. (B) Model of actin and myosin II stress fiber formation in human cardiomyocytes. Sarcomeres are templated by Muscle Stress Fibers (MSFs). MSFs do not require the Arp2/3 complex, and require the formin FHOD3. MSFs display slow retrograde flow compared with non-muscle stress fibers. Both NMIIA and NMIIB are localized to the edge of hiCMs, and display prominent NMII co-filaments. NMIIB-βCMII co-filaments are also present with MSFs. Large βCMII filament stacks form via concatenation and stitching of individual βCMII filaments.

https://doi.org/10.7554/eLife.42144.031

It has been well established that the actin filaments of actin arcs in non-muscle cells require both nucleation mediated by the Arp2/3 complex and formins (Henson et al., 2015; Hotulainen and Lappalainen, 2006; Murugesan et al., 2016). Our data would suggest the Arp2/3 is not formally required for MSF or sarcomere assembly in hiCMs. However, the Arp2/3 complex is localized to the edge of hiCMs, and future work will be needed to elucidate its role in cardiac biology. In contrast to the Arp2/3 complex, our data suggest that formins are required for MSFs and sarcomere assembly. Specifically, we found the formin FHOD3 has a major role in MSF and sarcomere assembly (Figure 12). This adds to the already established role of FHOD3 in sarcomere homeostasis (i.e., turn-over) (Iskratsch et al., 2010; Kan-O et al., 2012; Taniguchi et al., 2009). Future work will be required to elucidate the precise mechanism of how and where FHOD3 potentially nucleates actin filaments in sarcomeres. Canonical sarcomeric actin filaments have their barbed ends embedded within a Z-line (see schematic in Figure 1A). As such, this is where we would have predicted FHOD3 would localize. However, our SIM data show that FHOD3 does not localize to Z lines (Figure 6F, boxes 4, 5 and 6). Instead, FHOD3 appears to localize on either side of each Z-line. This could indicate that FHOD3 is only transiently associated with the barbed ends of canonical sarcomeric actin filaments. Alternatively, there could barbed ends of actin filaments in this region, which could indicate that the organization and/or dynamics of actin filaments within a sarcomere is more complex. Finally, while FHOD3 KD displayed the most severe phenotype, KD of DAAM1 and DIAPH1 in hiCMs resulted in both sarcomeric and non-sarcomeric defects in actin architecture. This agrees with previous literature showing roles for multiple formins in explanted mouse cardiomyocytes in maintenance of sarcomere structure (Rosado et al., 2014). The diverse roles of formins during sarcomere assembly and subsequent maintenance should be explored in future work.

We also show that NMIIA and NMIIB are required for proper sarcomere assembly in our hiCM model system. Interestingly, our data suggest that each motor may be playing a different role(s) in sarcomere assembly. NMIIB KD resulted in no detectable sarcomere assembly (Figure 8 and Figure 8—figure supplement 4), suggesting that NMIIB could be required for the initial and possibly subsequent steps of assembly. On the other hand, NMIIA KD hiCMs were able to assemble sarcomeres, although these were wispy with significantly shorter Z-lines (Figure 8 and Figure 8—figure supplement 3). This NMIIA KD-phenotype could be a result of several possibilities including a role of NMIIA in sarcomere maturation, alignment, or stability. Obviously, this is not an exhaustive list of potential mechanisms. In addition to showing NMIIA and NMIIB were required for sarcomere assembly, we show NMIIA and NMIIB form myosin II co-filaments with βCMII. βCMII filaments found in co-filaments were relatively small, (~300 nm), and subsequently grew larger as they transitioned to the larger filaments of the A-band (~1.6 microns), and lost NMII. Individual βCMII filaments (or a small bundle below the resolution limit of our imaging modality) concatenate to form the βCMII filament stacks of the A-band. This process requires the presence of actin tracks (Figure 12).

Collectively, our data provide new mechanistic and dynamic insight surrounding sarcomere assembly, and highlights key differences between classic, well-studied non-muscle stress fibers, and MSFs in cardiac myocytes (Figure 12). In addition, our model unifies certain aspects of previously proposed models of sarcomere assembly (Figure 12). These previously proposed models, while presented as mutually exclusive from one another, are actually quite similar in certain respects. Highlighting this, our data support a major feature shared between the Template Model and Pre-Myofibril Model. Specifically, that stress fibers are precursors to sarcomere containing myofibrils. These stress fibers were originally called ‘stress fiber-like structures’ when the Template Model was proposed (Dlugosz et al., 1984; Sanger et al., 2005). Later, they were renamed to Pre-Myofibrils in lieu of more thorough characterization (Rhee et al., 1994; Sanger et al., 2005). It was shown these stress fibers did not contain the same proteins as non-muscle stress fibers (Rhee et al., 1994). However, most of the known proteins in stress fibers found in cardiomyocytes have paralogs in non-muscle stress fibers. As such, we find ‘stress fiber-like structures’ an apt description. Therefore, we suggest simply calling these structures ‘stress fibers’ is preferable. In this study, we are comparing regulatory mechanisms of the non-muscle stress fiber referred to as actin arcs, to sarcomere precursors in cardiac myocytes. Thus, we decided to call them Muscle Stress Fibers (MSFs) to avoid confusion.

Our data support the concept that MSFs transition into sarcomere-containing myofibrils over time. This is the way the cartoon models of both the Template and Pre-Myofibril Models are presented (see original models (Dlugosz et al., 1984; Rhee et al., 1994)). An open question remains as to whether there is true templating during this process. The original Template Model posited that ‘stress fiber-like structures’ [MSFs] would disappear after they were used as a template to build a myofibril (Dlugosz et al., 1984). While some components of MSFs (e.g., α-actinin 2) do appear to persist during the MSF to sarcomere transition, others clearly do not (e.g., NMIIA/B). Furthermore, our data suggest that NMIIA/B filaments could be themselves a template for the addition of βCMII as all three paralogs can be found in co-filaments together in the region where NMIIA/B and βCMII overlap. In addition to the Templating/Pre-myofibril Models, our data also support aspects of the Stitching Model. The Stitching Model originally proposed that pre-assembled components of the sarcomere (i.e., A-bands and I-Z-I bodies) would stitch together sequentially to form a myofibril (Holtzer et al., 1997). We did not detect pre-formed I-Z-I bodies or A-bands in our assay (Figure 1—figure supplement 1) or sequential assembly of sarcomeres to form a myofibril (Figure 1). However, certain aspects of βCMII dynamics warrant comparison to the Stitching Model. We found separate βCMII filaments concatenated and ‘stitched’ together to form larger βCMII filament stacks (i.e., the A-band) (Figure 11C).

Our data show the transition of a MSF to a sarcomere-containing myofibril occurs on the dorsal (top) surface of the cell. However, a recent study also imaging iPSC derived cardiac myocytes claims that sarcomeres are formed on the ventral (bottom) surface of the cell near extracellular matrix adhesions (Chopra et al., 2018). This group also report that the first sarcomeres forms between 24– and 48 hr after plating, well after we detect the first sarcomeres appearing (Chopra et al., 2018). This led us to question what could be leading to these two seemingly opposite results. Importantly, it appears that this group was imaged the ventral (bottom) surface of their myocytes, as the focus of their study was on focal adhesions. Close inspection of their time-lapse movies revealed faint and blurred structures corresponding to sarcomeric patterns that show up in the frame right before the appearance of sarcomeres. This supports the notion that they are imaging sarcomeres that are coming into focus, and not assembling ‘de novo’. To test this idea, we imaged hiCMs with 3D confocal microscopy after they had been plated for 24 hr (Figure 12—figure supplement 1). While we also see similar patterns of sarcomeres appearing on the ventral surface as (Chopra et al., 2018), our data revealed these sarcomeres are moving down from the dorsal surface and not assembling on the ventral surface (Figure 12—figure supplement 1 and Video 5). Of interest, the phenomenon of actin arcs moving down to the ventral surface of non-muscle cells has also been reported previously (Gao et al., 2012; Hotulainen and Lappalainen, 2006). Finally, (Chopra et al., 2018) also claim that neither NMIIA nor NMIIB are required for sarcomere assembly. We also find this strange. While their double NMIIA/NMIIB knockout (KO) cardiomyocyte cell line has α-actinin 2 positive structures, they do not contain continuous labeled Z-lines aligned parallel to each other comparable to the control cell line. The authors did not measure Z-line lengths, spacing, or other criteria needed to define sarcomeres. These discrepancies between our study and theirs, including the role of NMII, need to be harmonized, as it will directly affect our interpretation of future in vivo data concerning sarcomere assembly.

Video 5
Sarcomeres traveling from dorsal to ventral surface of hiCM hiCM plated for 24 hr and expressing α-actinin 2-mCherry.

Four views from four Z-planes imaged using laser-scanning confocal microscopy. In the first frame, no sarcomeres can be visualized on the bottom of the cell (left most frame), but can clearly be seen in the 2 μm Z plane (middle right frame). As the video continues, the sarcomeres in the 2 μm Z plane travel towards the ventral (bottom Z plane) surface of the hiCM, which can be readily seen as the myofibril comes into focus first in the 1 μm Z plane (middle left), and then in the Bottom Z plane (left). Height: 38.2 μm. Video length: 17.5 hr.

https://doi.org/10.7554/eLife.42144.033

Data from in vivo studies attempting to answer the question of NMII contribution to sarcomere assembly in mice have been difficult to interpret (Sparrow and Schöck, 2009). Germline NMIIA KO mice fail to gastrulate and thus sarcomere assembly is impossible to examine (Conti et al., 2004). A conditional NMIIA KO mouse was generated using a promoter which is activated after heart formation has begun, and no heart defects were noted (Conti et al., 2015). A germline NMIIB KO mouse formed a functional heart, though most pups died before birth due to heart failure (Tullio et al., 1997). Only one high magnification image of the KO animal was shown, which demonstrated severe sarcomere disorganization (Tullio et al., 1997). Of interest, the NMIIB KO animals also showed highly increased NMIIA protein levels compared to controls (Tullio et al., 1997). Thus, the observed capacity of this animal to form sarcomere like structures could be due to genetic compensation by NMIIA. A conditional KO NMIIB mouse has also been made (Ma et al., 2009). While the authors showed an impressive NMIIB KD in the cerebellum via a neuronal specific driver, there were high levels of NMIIB protein in the heart at the time of analysis in the heart specific KO (Ma et al., 2009). In addition, the heart specific conditional KO is driven off of the alpha myosin heavy chain promoter, which switches on after sarcomere assembly has begun and indeed after the heart fields have begun to beat (Ma et al., 2009; Ng et al., 1991). Complicating the issue further, NMIIB has been difficult to localize in tissue. We believe this to be a result of paraffin embedding and subsequent paraffin removal and rehydration protocols. For example, we successfully localized NMIIB in formalin fixed human and mouse tissue (Figure 9 and Figure 9—figure supplement 1), but failed to localize NMIIB in paraffin embedded tissue. Future comparisons between in vivo and in vitro data sets will need to be addressed.

Intriguing and unanswered questions also remain to be tested. For example, how are the observed gradients of NMII and βCMII in cardiac myocytes established and maintained? Such questions have been previously difficult to impossible to test due complicated and technically challenging model systems. Here, we present a relatively easy to use model system to test these questions. The hiCMs we use in this study are commercially available. Our Methods outline protocols to transfect with DNA for protein expression and siRNA for protein knockdown, and trypsinization protocols for re-plating. Going forward, we believe studies using the experimental setup we describe here will not only continue to clarify previous models but also reveal new insights into both sarcomere assembly, and cardiac cell biology.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional
information
Cell line (human)iCell Cardiomyocytes^2Cellular Dynamics Internationalipsc derived cardiac myocytes
Cell line (human)Cor.4U CardiomyocytesNcardia (formerly Axiogenesis)ipsc derived cardiac myocytes
Cell line (human)HeLaAmerican Type Culture Collection (ATCC)CCL-2
Cell line (human)U2-OSAmerican Type Culture Collection (ATCC)HTB-96
Recombinant DNA reagentpHalo-NMHC IIA-C18this papersee Materials and methods for details on creation
Recombinant DNA reagentpHalo-NMHC IIB-C18this papersee Materias and methods for details on creation
Recombinant DNA reagentα-actinin-2-mEmeraldAddgene53988
Recombinant DNA reagentα-actinin-2-tdEOSAddgene57577
Recombinant DNA reagentNMIIA-(N-terminal)-mEGFPAddgene11347
Recombinant DNA reagentLifeact-mEmeraldGift from Michael Davidson
Recombinant DNA reagentLifeact-mAppleGift from Michael Davidson
Recombinant DNA reagentNMIIB-(N-terminal)-mEmeraldAddgene54192
Recombinant DNA reagentFHOD-mEGFP (lacking T(D/E)5XE exon)Gift from Dr. Elizabeth EhlerIskratsch et al. (2010)
Recombinant DNA reagentβCMII-mEGFPGenescript
(Piscataway, NJ, USA) this paper
see Materials and methods for details on creation
Recombinant DNA reagentp16b-eGFPGift from Dr. Jenniffer Lippincott-Schwartz and Dr. Pekka LappalainenLai, F. P., et al 2008.Koestler et al., 2013.
Antibodyrabbit polyclonal IgG anti-NMIIABiolegendP909801Used at 1:1000
Antibodyrabbit polyclonal IgG anti-NMIIBCell SignalingP3404SUsed at 1:200
Antibodyrabbit polyclonal
IgG anti-NMIIB
Biolegend909901Used at 1:200
Antibodymouse monoclonal IgG anti-βCMIIIowa Hybridoma BankA4.1025Used at 1:2
Antibodyrabbit polyclonal IgG anti-α-actinin-2Sigma-Aldrichclone EA-53Used at 1:200
Antibodyrabbit polyclonal IgG anti-p34-Arc/ArpC2 (Arp2/3 complex)Millipore Sigma07–227Used at 1:100
Antibodyrabbit polyclonal IgG anti-DAAM1Bethyl LaboratoriesHPA026605Used at 1:100
Antibodyrabbit polyclonal IgG anti-DIAPH1Sigma-AldrichA300-078AUsed at 1:100
Antibodymouse monoclonal IgGanti-cardiac troponin TSanta Cruz BiotechnologyCT3, sc-20025Used at 1:50
Antibodygoat anti mouse IgG Alexa Fluor 568ThermoFisher ScientificA-11004Used at 1:100
Antibodygoat anti rabbit IgG Alexa Fluor 568ThermoFisher ScientificA-11011Used at 1:100
Antibodygoat anti mouse IgG Alexa Fluor 488ThermoFisher ScientificAA28175Used at 1:100
Antibodygoat anti rabbit IgG Alexa Fluor 647ThermoFisher ScientificA-21244Used at 1:100
Antibodygoat anti mouse IgM Alexa Fluor 488ThermoFisher ScientificA-21042Used at 1:100
Antibodymouse monoclonal IgG anti-FHOD3Santa Cruz BiotechnologyG-5, sc-374601Used at 1:100
Sequence-based reagentSmartPool siRNA DIAPH1GE DharmaconE-010347-00-0005
Sequence-based reagentSmartPool siRNA DAAM1GE DharmaconE-012925-00-0005
Sequence-based reagentSmartPool siRNA FHOD3GE DharmaconE-023411-00-0005
Sequence-based reagentSmartPool siRNA MYH10 (NMIIB)GE DharmaconE-023017-00-0010
Sequence-based reagentSmartPool siRNA MYH9 (NMIIA)GE DharmaconE-007668-00-0005
Commercial assay or kitRneasy Mini KitQiagen74104
Commercial assay or kitQIAprep spin Miniprep Kit (250)Qiagen27106
Chemical compound, drugBlebbistatinSigma-AldrichB0560
Chemical compound, drugLatrunculinBSigma-AldrichL5288
Chemical compound, drugLatrucluinASigma-AldrichL5163
Chemical compound, drugCK666Sigma-AldrichSML0006
Chemical compound, drugSMIFH2Sigma-AldrichS4826
Chemical compound, drugTransIT-TKOMirus BioMIR 2150
Chemical compound, drugFuGENE HDPromegaE2311
Chemical compound, drugAlexa Fluor 488 PhalloidinThermoFisher ScientificA12379
Chemical compound, drugViafectpromegaE4981
Software, algorithm(Fiji Is Just) ImageJNIH (open source)

Contact for reagent and resource sharing

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Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, D.T.B. (dylan.burnette@vanderbilt.edu)

Experimental model and subject details

Cell line growth and experimental conditions

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Human induced pluripotent stem cell cardiomyocytes (hiCMs) were purchased from either Axiogenesis (Cor.4u, Ncardia Cologne, Germany) or Cellular Dynamics International (iCell caridomyocytes2, Madison, WI). Cells were cultured as per manufacturer’s instructions. hiCMs were cultured in 96 well plates and maintained in proprietary manufacturer provided media until ready for experimental use. Knockdown experiments in hiCMs were started 4 days after the initial thaw. See Plating Assay for more detailed protocol for cell plating. See below for detailed information on protein transfection and siRNA mediated knockdown

U-2 OS and HeLa cells (HTB-96; American Type Culture Collection, Manassas, VA) cells were cultured in 25 cm2 cell culture flasks (25-207; Genessee Scientific Corporation, San Diego, CA) with growth medium comprising DMEM (10–013-CV; Mediatech, Manassas, VA) containing 4.5 g/L L-glutamine, D-glucose, and sodium pyruvate and supplemented with 10% fetal bovine serum (F2442; Sigma-Aldrich, St. Louis, MO). For protein expression experiments in U-2 OS cells, cells were transiently transfected with FuGENE 6 (E2691; Promega, Madison, WI) according to the manufacturer’s instructions. Knockdown of NMIIA and NMIIB was performed as previously, with the Accell SMARTpool siRNA to human MYH9, MYH10 or Accell scrambled control purchased from Thermo Fisher Scientific (Waltham, MA) in combination with Lipofectamine 2000 Transfection Reagent (cat# 11668027, Thermo Fisher Scientific, Waltham, MA) for HeLa cells. For both live and fixed cell microscopy, cells were plated and imaged on 35 mm glass bottom dishes with a 10 mm micro-well #1.5 cover glass (Cellvis, Mountain View, CA) coated with 25 µg/mL laminin (114956-81-9, Sigma-Aldrich).

Plasmids

Plasmids encoding NMIIA-(N-terminal)-mEGFP (11347; Addgene, Cambridge, MA) with mEGFP on the N-terminus of NMIIA heavy chain were used as described previously (Chua et al., 2009). Plasmid encoding Lifeact-mEmerald and Lifeact-mApple were gifts from Michael Davidson. Plasmid encoding NMIIB-(N-terminal)-mEmerald was purchased from Addgene (54192; Addgene, Cambridge, MA). The plasmid encoding α-actinin 2-tdEOS was purchased from Addgene (57577; Addgene, Cambridge, MA). Plasmid encoding human βCMII was synthesized by Genscript (Piscataway, NJ, USA). Briefly, the wild-type human MYH7 (βCMII) sequence from the National Center for Biotechnology Information (NCBI, Bethesda, MD, USA) was cloned into a pUC57 along with Gateway DNA recombination sequences in order to facilitate rapid fluorescent protein integration and swapping (Gateway Technology, ThermoFisher Scientific, Waltham, MA). mEGFP containing a previously published linker sequence was added to the βCMII plasmid using Gateway Vector Conversion System with One Shot ccdB Survival Cells (ThermoFisher Scientific, Waltham, MA) for the βCMII-(N-terminal)-mEGFP construct used in this study. FHOD3-mEGFP plasmid lacking the T(D/E)5XE exon was a gift from Elizabeth Ehler. This construct has previously been shown to localize to sarcomeres in neonatal rat cardiomyocytes (Iskratsch et al., 2010) (see top left panel of Figure 2A in reference). Assembling pHalo-NMHC IIB-C18: pEmerald-NMHC IIB-C18 (Addgene, #54192) was the kind gift of Michael Davidson. Halo tag cDNA was PCR amplified from pHalo-N1 and subcloned into the Age I/BspEI mEmerald site of pEmerald-NMHC IIB-C18, replacing mEmerald, to generate the sequence verified pHalo-NMHC IIB-C18 construct. Assembling pHalo-NMHC IIA-C18 (NMIIA-HaLo): The NheI/BspEI Halo cDNA fragment from the pHalo-NMHC IIB-C18 construct was subcloned into the NheI/BspEI mEmerald site of pEmerald-NMHC IIA-C18 (Addgene, #54190) (Burnette et al., 2014), replacing mEmerald, to generate the sequence verified pHalo-NMHC IIA-C18 construct. The pEGFP-p16b plasmid (to visualize the Arp2/3 complex) was a gift from Dr. Pekka Lappalainen and previously published (Koestler et al., 2013; Lai et al., 2008).

Halo-tag Labeling

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hiCM expressing Halo-tag constructs were labeled with JF Dye 585 at 1:100 dilution in pre-warmed culture medium for 1 hr. The plate was then washed by gently exchanging with fresh medium 3 times (Grimm et al., 2017; Grimm et al., 2015)

Cell line authentication

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The HeLa cell line used in this study was a gift of Dr. DA Weitz (Harvard University). The Burnette lab had this line authenticated by Promega and ATCC using their ‘Cell Line Authentication Service’ in 2015. The methods and test results received from Promega and ATCC are as follows:

“Methodology: Seventeen short tandem repeat (STR) loci plus the gender determining locus, Amelogenin, were amplified using the commercially available PowerPlex 18D Kit from Promega. The cell line sample was processed using the ABI Prism 3500xl Genetic Analyzer. Data were analyzed using GeneMapper ID-X v1.2 software (Applied Biosystems). Appropriate positive and negative controls were run and confirmed for each sample submitted.’

“Data Interpretation: Cell lines were authenticated using Short Tandem Repeat (STR) analysis as described in 2012 in ANSI Standard (ASN-0002) Authentication of Human Cell Lines: Standardization of STR Profiling by the ATCC Standards Development Organization (SDO)’

“Test Results: The submitted profile is an exact match for the following ATCC human cell line(s) in the ATCC STR database (eight core loci plus Amelogenin):CCL-2 (HeLa)’

The U2 OS cell line used in this study was purchased directly from ATCC by the Burnette lab in 2014 (ATCC, HTB-96). Mycoplasma Monitoring: Both HeLa and U2 OS cell lines were checked for potential mycoplasma infection using either DAPI or Heoscht throughout the course of this study.

Live-Cell sarcomere assembly visualization assay

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To visualize sarcomere formation, we developed a repeatable method, which can be used to visualize any fluorescently tagged protein during sarcomere formation in hiCMs. Prior to performing this assay, cells are maintained in desired culture vessel (our hiCMs were maintained in 96 well plates). In this assay, cells are transiently transfected with desired fluorescently tagged protein, trypsinized, plated on desired imaging dish, and imaged to observe sarcomere formation. A sarcomere assembly assay proceeds as follows. First, hiCMs are transfected with desired fluorescently tagged protein as described below (Viafect, overnight transfection). Transfection mix is washed out with culture media. Cells are then detached from culture vessel using trypsinization method described below. Cells are then plated on a desired culture vessel (in this study, 10 mm #1.5 glass bottom dishes were used, CellVis, Mountain View, CA) for live cell imaging. Imaging vessels were pre-coated with 25 µg/mL laminin for 1 hr at 37° C and washed with 1x PBS containing no Mg2+/Ca2+. Cells were allowed to attach for ~1.5 hr and media was added to fill the glass bottom dish. Cells were then imaged using desired imaging modality at 3–6 hr post plating. The 3–6 hr time window was optimal, as cells had not yet established sarcomeres, but were healthy enough to tolerate fluorescence imaging. This powerful assay can be adapted to visualize the kinetics during sarcomere formation of any fluorescently tagged protein. We have used this assay to visualize actin (Lifeact), FHOD3, βCMII, α-actinin 2, NMIIA, and NMIIB. The latter two proteins being large (i.e.,>250 KD).

Trypsinization of hiCMs

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To trypsinize Cellular Dynamics hiCMs, manufacturers recommendations were used, as follows. All volumes apply were modified from 24 well format for 96 well plates. hiCMs were washed 2x with 100 uL 1x PBS with no Ca2+/Mg2+ (PBS*). PBS* was completely removed from hiCMs and 40 uL 0.1% Trypsin-EDTA with no phenol red (Invitrogen) was added to hiCMs and placed at 37° C for 2 min. Following incubation, culture dish was washed 3x with trypsin inside well, rotated 180 degrees, and washed another 3x. Trypsinization was then quenched by adding 160 µL of culture media and total cell mixture was placed into a 1.5 mL Eppendorf tube. Cells were spun at 1000gs for 5 min, and supernatant was aspirated. Cells were then re-suspended in 200 uL of culture media and plated on a 10 mm glass bottom dish pre-coated with 25 µg/mL laminin for 1 hr. Cells were then allowed to attach for at least 1 hr, and 2–3 mLs of culture media with or without drug was added to cells.

To trypsinize Axiogenesis hiCMs, manufacturers recommendations were used, as follows. Cells were washed 2x with 500 µL PBS*. Cells were placed in 37° C incubator for 7 min in PBS*. Following 7 min, PBS* was aspirated and 40 µL 0.5% Trypsin (Invitrogen) was placed on cells for 3 min in 37° C incubator. Following 3 min, 160 µL full Cor.4u media was used to quench trypsinization and re-suspend cells. Cells were then plated on pre-coated glass bottom dish and media added 1.5 hr later to dilute trypsin and fill chamber. Note* this trypsinization protocol has since been modified by Axiogenesis (now Ncardia). See manufacturer for new protocol.

It is important to note that the trypsinization protocol is based off of the hiCMs manufacturers protocols for cell plating, and these hiCMs have been trypsinized prior to functional assays of cardiomyocyte performance and characterization (e.g., drug response, electro-physiology, maturation, etc…) (Fine et al., 2013; Ivashchenko et al., 2013; Mioulane et al., 2012).

Transient transfection of hiCMs

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Cellular Dynamics hiCMs were transfected via modification of manufacturer’s recommendations as follows. Volumes used are for transfection in 96 well plates. 2 µL of total 200 ng plasmid (containing fluorescently tagged protein of interest, diluted in Opti-MEM) and 2 µL 1:5 diluted Viafect (Promega, E4981, in Opti-MEM) was added to 6 µL Opti-MEM. Entire mixture of 10 µL was added to single 96 well of hiCMs containing freshly exchanged 100 µL full culture media. Transfection was allowed to go overnight (~15 hr), and washed 2x with full culture media. For transfection of multiple probes, 2 µL of 200 ng plasmid was used for each probe together with 4 µL 1:5 diluted Viafect, into 2 µL Opti-MEM and mixture was applied to cells as above.

Axiogenesis hiCMs were transfected via modification of manufacturer’s recommendations as follows. A 3.5:1 Fugene to DNA ratio was used to transfect Axiogenesis hiCMs. 1.2 µL Fugene +0.33 µg DNA per 96 well into 5 µL serum free Cor.4u media was incubated for 15 min at room temperature. 95 µL full Cor.4u media was added to mixture and entire mixture was added to hiCMs (on top of 100 µL already in well). For transfection of 2 separate plasmids, 3.5:1 ratio was used for both plasmids and additional volume was subtracted from 95 µL dilution. Note* this transfection protocol has since been modified by Axiogenesis. See manufacturer for new protocol.

Protein knockdown

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Cellular Dynamics hiCMs were used for knockdown experiments via modification of manufacturers recommendations. Volumes used are for siRNA application in 96 well plates. Dharmacon SmartPool siRNA (GE Dharmacon, Lafayette, CO) targeted to MYH9 (NMIIA), MYH10 (NMIIB), FHOD3, DAAM1, and DIAPH1 were used (E-007668-00-0005, E-023017-00-0010, E-023411-00-0005, E-012925-00-0005, and E-010347-00-0005, respectively). To achieve KD, a master mixture of 100 μl Opti-MEM (ThermoFisher, Waltham, MA)+4 μl Transkit-TKO (Mirus Bio, Madison WI)+5.5 μl 10 μM siRNA was incubated for 30 min at room temperature. 80 μl of fresh, pre-warmed media was added to hiCMs. Following incubation of siRNA mixture, 8.3 μl of mixture was added to each individual well of 96 well plate. hiCMs were then incubated for 2 days at 37° C. hiCMs were then washed 2x with fresh, pre-warmed media. To achieve KD of NMIIA, NMIIB, FHOD3, DAAM1, and DIAPH1, 3 rounds of siRNA mediated KD described above were necessary. Following 3 rounds of scramble control siRNA treatment, hiCMs still beat and maintained sarcomere structure (see Figure 8—figure supplement 6).

Pharmacological experiments

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For all pharmacological experiments, drugs were added to pre-warmed and equilibrated media before adding to hiCMs. For sarcomere assembly assays, 25 μM SMIFH2 (Sigma-Aldrich, S4826), 25 μM CK666 (Sigma-Aldrich, SML0006), and 100 μM Blebbistatin (Sigma-Aldrich, B0560) (separately) were added 1.5 hr after plating to allow hiCMs to attach to the substrate. hiCMs were subsequently fixed (see below) at 24 hr post plating. For live-cell experiments (as in Figure 5D), media in imaging container was aspirated using a vacuum system, and media containing drug was added to hiCMs on the microscope stage. This allowed the same hiCM to be imaged pre and post-drug application. For the LatrunculinB experiment (Figure 9J), cells were allowed to spread for 18 hr in normal media, and media containing 5 μM LatrunculinB (Sigma-Aldrich, L5288) was added for 6 hr and hiCMs were fixed (for a total spreading time of 24 hr).

Western blotting

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Gel samples were prepared by mixing cell lysates with LDS sample buffer (Life Technologies, #NP0007) and Sample Reducing Buffer (Life Technologies, #NP00009) and boiled at 95°C for 5 min. Samples were resolved on Bolt 4–12% gradient Bis-Tris gels (Life Technologies, #NW04120BOX). Protein bands were blotted onto a nylon membrane (Millipore). Blots were blocked using 5% NFDM (Research Products International Corp, Mt. Prospect, IL, #33368) in TBST. Antibody incubations were also performed in 5% NFDM in TBST. Blots were developed using the Immobilon Chemiluminescence Kit (Millipore, #WBKLS0500).

Fixation and immunohistochemistry

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hiCMs and HeLa cells were fixed with 4% paraformaldehyde (PFA) in PBS at room temperature for 20 min and then extracted for 5 min with 1% Triton X-100% and 4% PFA in PBS as previously described (Burnette et al., 2014). Cells were washed three times in 1 × PBS. To visualize endogenous βCMII and transfected proteins in fixed cells, hiCMs were live-cell extracted before fixation as described previously to reduce background and non-cytoskeletal myosin II filaments (i.e., the soluble pool). Briefly, a cytoskeleton-stabilizing live-cell extraction buffer was made fresh containing 2 ml of stock solution (500 mM 1,4-piperazinediethanesulfonic acid, 25 mM ethylene glycol tetraacetic acid, 25 mM MgCl2), 4 ml of 10% polyoxyethylene glycol (PEG; 35,000 molecular weight), 4 ml H2O, and 100 μl of Triton X-100, 10 μM paclitaxel, and 10 μM phalloidin. Cells were treated with this extraction buffer for 1 min, followed by a 1 min wash with wash buffer (extraction buffer without PEG or Triton X-100). Cells were then fixed with 4% PFA for 20 min. After fixation, the following labeling procedures were used: for actin visualization, phalloidin-488 in 1 × PBS (15 μl of stock phalloidin per 200 μl of PBS) was used for 3 hr at room temperature. For immunofluorescence experiments, cells were blocked in 10% bovine serum albumin (BSA) in PBS. Primary antibodies were diluted in 10% BSA. NMIIA antibody (BioLegend, P909801) was used at 1:1000. NMIIB antibody (Cell Signaling, 3404S and BioLegend 909901) were used at 1:200. βCMII antibody (Iowa Hybridoma Bank, A4.1025) was used undiluted from serum. α-actinin 2 (Sigma-Aldrich, clone EA-53) antibody was used at 1:200. FHOD3 antibody (Santa Cruz Biotechnology, G-5, sc-374601) was used at 1:100. Arp2/3 antibody (Anti-p34-Arc/ARPC2, Millipore Sigma, 07–227) was used at 1:100. DAAM1 antibody (Bethyl Laboratories, A300-078A) was used at 1:100. DIAPH1 antibody (Sigma-Aldrich, HPA026605) was used at 1:100. Cardiac Troponin T antibody (Santa Cruz Biotechnology, CT3, sc-20025) was used at 200 µg/ml. Secondary antibodies were diluted in 10% BSA at 1:100 and centrifuged at 13,000 rpm for 10 min before use. Cells were imaged in VECTASHIELD Antifade Mounting Media with DAPI (H-1200, VECTOR LABORATORIES, Burlingame, CA, USA). To label both NMIIA and NMIIB in the same sample (as in Figure 7A), NMIIA primary antibody was directly labeled using a primary antibody labeling kit from Biotium (Mix-n-Stain CF Antibody Labeling Kits, Biotium, Inc. Fremont, CA). Following manufacturers protocol, NMIIA was primary labeled, and stain was visually compared to standard immunofluorescence protocol (see above) to validate localization pattern.

RNA-seq data analysis

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RNA-seq reads were aligned to the human reference genome hg19 using STAR (Dobin et al., 2013) and quantified by featureCounts (Liao et al., 2014). Read counts were normalized by the Relative Log Expression (RLE) method and FPKM values for each sample were generated by DESeq2 (Love et al., 2014).

Structured illumination microscopy

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Two SIM microscopes were used in this study (individual experiments were always imaged using the same microscope). SIM imaging and processing was performed on a GE Healthcare DeltaVision OMX equipped with a 60×/1.42 NA oil objective and sCMOS camera at room temperature. SIM imaging and processing was also performed on a Nikon N-SIM structured illumination platform equipped with an Andor DU-897 EMCCD camera and a SR Apo TIRF (oil) 100 × 1.49 NA WD 0.12 objective at room temperature.

Spinning disk microscopy

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Spinning disk confocal images were taken on a Nikon Spinning Disk equipped with a Yokogawa CSU-X1 spinning disk head, Andor DU-897 EMCCD camera and 100x Apo TIRF (oil) 1.49 NA WD 0.12 mm objective at 37 degrees C and 5%CO2.

Laser-scanning confocal microscopy

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Laser-scanning confocal images were taken on a Nikon A1R laser scanning equipped with a 60x/1.40 Plan Apo Oil objective at 37 degrees C and 5%CO2. Confocal images for Figure 9C (in vivo, right) and Figure 9—figure supplement 1C-D were taken on a Zeiss 880 with AiryScan equipped with a 63x/1.40 Plan-Apochromat Oil objective at room temperature.

Wide-field microscopy

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Wide-field images were taken on a high-resolution wide-field Nikon Eclipse Ti equipped with a Nikon 100x Plan Apo 1.45 NA oil objective and a Nikon DS-Qi2 CMOS camera. Data presented in Figure 8—figure supplement 6 was acquired on an Essen Incucyte S3 (Ann Arbor, MI) Live-Cell Analysis system with the 20x objective. Photo-conversion experiments were performed as previously described (Fenix et al., 2016).

Quantification and statistical analysis

Sarcomere assembly quantification

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To quantify percent of hiCMs with sarcomeres (i.e., Figure 4B), the actin cytoskeleton (via fluorescently labeled phalloidin) was imaged using structured illumination microscopy. hiCMs were quantified as containing sarcomere structures if they contained at least one myofibril containing at least 3 Z-discs (bright phalloidin staining which overlaps with α-actinin 2 staining as in Figure 4A) spaced ~1.5–2 μm apart. By these metrics our quantification of sarcomere formation is not a measure of sarcomere maturity or alignment, but a measure of the hiCMs ability to assemble the building blocks of the sarcomere (i.e., the thin actin filaments) in response to a perturbation. Thus, while NMIIA KD hiCMs clearly form unaligned, disorganized, and fewer sarcomeres and myofibrils than control hiCMs, they still maintain the ability to assemble sarcomere structures, which is reflected in our quantification (Figure 8A and C). In the same vein, we realize our quantification is a very liberal quantification of sarcomere assembly. While we don’t expect a small array of sarcomeres to represent a functionally capable cardiomyocyte, we are investigating the early steps of sarcomere assembly and the ability of hiCMs to form the basic actin-myosin structure of the sarcomere.

β cardiac myosin II (βCMII) filament and stack quantification

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A similar methodology was used to quantify βCMII A-band filament stacks using endogenous βCMII staining and SIM instead of the actin cytoskeleton. A βCMII filament was quantified as the minimum SIM resolvable βCMII unit which had a bipolar organization (a filament with motor domains on each side), as represented in Figure 11A. hiCMs were quantified as containing βCMII A-band filament stacks if they contained even one βCMII filament stack in the cell. A βCMII filament stack was defined as being thicker than the minimum SIM resolvable βCMII filament, indicating multiple SIM resolvable βCMII filaments. Indeed, by this metric, βCMII filament stacks have more resolvable ‘motor-domains’ than βCMII filaments.

For actin arc and MSF retrograde flow rates (as in Figure 3), 3 regions of interest (ROIs) were used per cell. ROIs were drawn using the line tool in FIJI starting from in front of the leading edge (to ensure new MSF formation was captured) to the cell body where sarcomere structures were localized. ROIs were then used to measure MSF translocation rates using the kymograph tool (line width = 3) on hiCMs which had been aligned using StackReg function. Kymographs generated in this manner were then manually measured by counting pixels on the X axis (distance) and the Y axis (time) for a distance/time measurement resulting in translocation rates. This method is similar to previously described methods of actin arc translocation in non-muscle cells.

To measure distance between α-actinin 2 structures, the line tool and measure tool in FIJI (Fiji is Just ImageJ) was used. Lines were drawn, positions recorded (using ROI tool), and distances measured between α-actinin 2 structures. Multiple regions per cell for α-actinin 2 in MSFs, and whole cells for distances between sarcomeres were used.

To measure Z-line sizes in hiCMs, the line tool and measure tool in FIJI was used. Lines were drawn on individual Z-lines and their positions were recorded using the ROI tool. This gave a measurement of Z-line size and position within the cell. For Z-line measurements, such as Figure 8E, all Z-lines which could be reliably measured were measured.

Immunofluorescence localization quantification

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To quantify localization of NMIIA, NMIIB, and βCMII, line scans starting from the edge of the cell were taken for every cell and the normalized average localizations were used to average the number of indicated experiments for the final localization patterns depicted in the graph (as in Figure 7B).

To quantify Arp2/3 intensity, a similar but altered strategy was taken. four separate boxes for each cell were placed along the edge of the cell (i.e., the lamellipodium) using the actin channel for guidance. These boxes were then used to measure average intensity of the anti-p34 channel (i.e., the Arp2/3 complex). Background subtracted averages for each cell in control and CK666 treated hiCMs were used to quantify percent decrease as depicted in Figure 4F.

Cells used for this study

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In order to have comparable results, cells used for this study were standardized based on morphology. Specifically, spread non-muscle cells and hiCMs with a broad leading edge and lamella were chosen as previously shown in studies of both non-muscle and muscle contractile system formation (Burnette et al., 2014; Rhee et al., 1994). This also facilitated the ability to observe the MSF to sarcomere transition in live hiCMs, and is recommended for future studies investigating sarcomere assembly. All experiments measuring sarcomere assembly were conducted at least 3 times, separately, and cells were imaged using SIM. Though this results in a relatively small number of cells for some of the experiments, we believe super-resolution imaging modalities such as SIM offer invaluable insight into sarcomere assembly (Gustafsson, 2005). Indeed, sub-diffraction imaging is required to reliably localize myosin II co-filaments both in vitro and in vivo. Furthermore, as has been seen in Drosophila, even high-resolution imaging modalities such as laser-scanning confocal microscopy are not sufficient to detect subtle, yet important, structural changes in response to perturbation (Fernandes and Schöck, 2014).

Statistics

Statistical significance was calculated using 2-tailed, unpaired Students T-tests performed in Excel. All independent experiments represent biological replicates. Error bars in all graphs represent standard error of the mean (SEM). For all graphs depicting % of cells (for example, Figure 1D), number of cells and experiments is indicated in figure legend. Percents displayed represent the average of the averages of the all experiments performed. For example, if controls cells displayed 95%, 90%, and 100% of all cells displaying actin arcs in three separate experiments, the represented percentage in the graph would be 95%. SEM was calculated by dividing the standard deviation by the square root of the number of separate experiments. For actin translocation rates at least three measurements per cell were used to calculate the average translocation rates per cell. Translocation rates were then calculated in same manner as described above. Line-scan graphs represent normalized relative fluorescence.

References

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Decision letter

  1. Anna Akhmanova
    Senior and Reviewing Editor; Utrecht University, Netherlands

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: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for submitting your work entitled "Muscle stress fibers are essential sarcomere precursors and are regulated differently than non-muscle stress fibers" for consideration by eLife. Your article has been reviewed by a Senior Editor, a Reviewing Editor, and three reviewers. The reviewers have opted to remain anonymous.

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 this paper, the authors investigated sarcomere assembly in human iPSC- derived cardiomyocytes and provide novel molecular insights into this process, which are potentially of broad interest. However, all three reviewers felt that there is a very significant number of issues that need to be addressed before this paper can be published. In particular, the referees found that some data were incomplete and some controls missing, that important quantifications were lacking, and that certain data needed to be analyzed differently. Furthermore, a number of points had been raised that would require addition of new data, such as evaluation of the spacings and dynamics of α-actinin puncta, two-color imaging of myosin II and β cardiac myosin II, and photoactivation experiments to link muscle stress fibers to sarcomere assembly more convincingly. We return the paper to you, hoping that you will find the detailed comments of the reviewers useful for improving your study. However, we will be prepared to consider a new submission, which would fully address the comments of all three reviewers. In case you decide to resubmit your paper to eLife, it will be sent to the same Reviewing Editor, who will then decide, based on your revisions, whether the paper warrants re-review and whether the same reviewers should be consulted.

Reviewer #1:

In this manuscript Fenix et al., have reported a novel phenomenon in sarcomere assembly in human IPS derived cardiomyocytes (hiCM), which builds on prior knowledge in the field. Using confocal, super-resolution and live cell imaging assays in hiCM, the authors demonstrate that sarcomeres assemble from F-actin stress fibers organized in arcs near the edge of the spreading cells, that they term muscle stress fibers (MSF). These resemble actin arcs in spreading cells from HeLa and U2OS cell lines but have quantitative differences. They showed that retrograde movement of the MSFs results in maturation of sarcomeres (ie formation of striated myofibrils) and that the rate of retrograde flow of MSFs is significantly slower than that of F-actin arcs in the non-muscle cell types. These observations of myofibril assembly in the hiCMs are overall similar to a number of previous studies using fluorescent-tagged α-actinin to image the transitions of stress fiber-like structures (SFLSs)/pre-myofibrils into myofibrils in primary cardiac myocytes, but numbers and rates have now been quantified here (note the terminology is also different here). In a new advance, they go onto show that the formin FHOD3 rather than Arp2/3 plays pivotal roles in MSF directed myofibril assembly/sarcomere formation (by contrast, Arp2/3 is important for assembly of non-muscle cell F-actin arcs). In addition, they demonstrate novel functions for NMIIA versus NMIIA in myofibril assembly, showing that in the MSF to sarcomere transformation process NMIIB is necessary, while NMIIA appears to play a redundant role, using an acute knockdown approach. Finally, the mechanism of β-cardiac myosin II thick filament assembly into A bands was also studied, and live cell imaging was used to evaluate the mechanism of thick filament assembly into sarcomeres in striated myofibrils. Through these experiments Fenix et al., have convincingly shown that MSFs (which likely correspond to previously-termed SFLSs or pre-myofibrils) are precursors for maturing sarcomeres and require a different non-muscle myosin isoform as compared to the arc-like stress fibers found in non-muscle cell lines. However, some data are weak and do not add to the main points (e.g., the zebrafish data), and there are some missing pieces in the data (e.g., lack of direct comparisons to the previous literature using α-actinin2 as a marker for myofibril assembly, questions regarding location of FHOD3, potential myosin isoform compensation), that need to be addressed to place this study in the context of previous work. Many of the figures also need to be improved to strengthen the points and clarify some ambiguous results.

Main points:

1) The authors claim in the Introduction that the hiCM system allows them to study de novo myofibril assembly in vitro for the first time, unlike the previous primary cardiac cell cultures from embryonic chick hearts or neonatal mouse hearts. However, Figure 5 shows that the hiCM cells contain sarcomeres before replating and spreading! Thus, the hiCM system is actually quite similar to the primary cell systems, which disassemble and then reassemble sarcomeres after replating and spreading. Nevertheless, the hiCM cells are a great system, and the authors present some interesting and novel findings regarding the roles of actin arcs and myosin isoforms in myofibril assembly. The Introduction and Results section should be rewritten and shortened to emphasize the new work on formins and myosin isoforms.

2) There is another very important point regarding previous work on myofibril assembly. The authors state that they are unable to find I-Z-I bodies in the hiCMs, but I can clearly see small α-actinin puncta along the F-actin arcs in Figure 2A. The authors may have been misled by the vast and confusing literature; I-Z-I bodies are simply small α-actinin2 puncta linearly arranged along F-actin bundles that stain continuously for F-actin. These do appear to be present in the hiCMs. The authors should reexamine their control (untreated) hiCMs and present some high mag images of the cell edge and the F-actin arcs stained for α-actinin2. How does the spacing of α-actinin2 puncta in the arcs/MSFs of hiCMs compare with the literature on myofibril assembly in other cardiac cell types? The distances between the α-actinin2 puncta should be measured in the various experiments, including the formin and myosin KD experiments. It is important to compare the so-called MSFs in this study with the various so-called "SFLSs" and "pre-myofibril" etc structures studied by other groups. Most of the live imaging in previous work used fluorescent-α-actinin2 to study formation of myofibrils; thus, one would like to know whether this study obtains similar results. For example, closely-spaced α-actinin puncta can be seen transitioning into the wider striations in the myofibrils as they move inwards at the spreading edge of a chick cardiac myocyte in Figure 5 in Dabiri et al., 1997. I think these could be the arcs that are shown here in the hiCMs. There are other types of examples in the literature.

3) As a related point, what is the direct evidence that the F-actin arcs in the hiCMs are similar to the F-actin arcs in non-muscle cells? The first paragraph of the Results section states: "Strikingly, super-resolution imaging revealed the MSFs in hiCMs resembled a classic actin stress fiber found in non-muscle cells, referred to as actin arcs (Figure 1C and 1D)." I expected the figure would show high mag images of the hiCM arc substructure compared to the non-muscle cell arcs, but this figure shows relatively low mag images of the arcs in U2OS and HeLa cells, revealing continuous F-actin along the fibers in all cell types. How does this show that the MSFs are similar to the arcs in the non-muscle cells? We need to see some high-resolution images of the arcs in the hiCMs and the non-muscle cells – for example what are the spacings between the Z bodies (α-actinin puncta) in the arcs in each cell type? I would like to know whether the NMIIA and NMIIB are located in between the α-actinin2 puncta of the MSFs, which would also be similar to the SFs in the non-muscle cell types (but with a different α-actinin isoform).

4) Actin arcs are shown in the live cell imaging experiments to be precursors for "myofibrils containing sarcomeres" (note- this is a more precise terminology- a sarcomere is a small unit of a myofibril- and the arc may have a small actomyosin 'mini-sarcomere' type structure similar to the stress fibers of non-muscle cells). Can other types of actin stress fibers like dorsal and ventral stress fibers also mature into sarcomeres in hiCM? Or, do these hiCM cells only possess actin arcs? This seems unlikely, as I think I can see some radial stress fibers in the cells in Figure 3E and in Figure 1H near the edge.

5) The authors have shown that FHOD3 knockdown in hiCM cells results in reduced sarcomere formation. However, they have not shown the cellular distribution of FHOD3 by immunofluorescence, unlike Arp2/3. Where does FHOD3 localize in the hiCM in the initial MSF stage, and later when the sarcomeres are formed? Does FHOD3 directly interact with the MSFs and/or the sarcomeres?

6) The authors use a pan-formin inhibitor, SMIFH2, to show that formins are important for formation of the actin arcs and sarcomeres. While FHOD3 is clearly important, their experiments do not rule out roles for other formins. For example, see Rosado et al., 2014. Also, the authors claim that mDia1 is not involved in the MSFs, but this has not been tested here. This could be tested, or at the least, discussed and clarified.

7) NMIIA and NMIIB can form heteropolymers in hiCM, yet NMIIB knockdown alone disrupts sarcomere formation. Is this because there is a compensatory rise in NMIIB expression upon NMIIA knockdown, which restores the total NMII levels in the cell? However, another possibility is that the NMIIA levels are not enhanced upon NMIIB knockdown, resulting in a drop of cellular total NMII levels, bringing about a disruption of sarcomere assembly. The authors should examine the NMIIA levels upon NMIIB knockdown, and vice versa, to address this issue. It would also be helpful to provide a quantitative estimate of NMIIA/NMIIB distribution in the soluble and insoluble fractions of hiCM cells during the MSF and sarcomere stages. This may confirm why one isoform is more important than the other in sarcomere formation.

8) Although the results show that NMIIB is essential for de-novo sarcomere assembly in spreading cells, is it also important for the maintenance of the sarcomeres?

9) I appreciate the authors' efforts to study NMIIB in vivo, but the experiments are not convincing. The zebrafish morpholino experiments are potentially exciting but are underdeveloped and missing important controls. What is the result of MO for MYH9 (NMIIA)? MO of the MYH10 (NMIIB) could also affect heart development via effects on non-cardiac cell types, e.g., vascular endothelium. A rescue using a cardiac-specific MYH10 transgene would address this. Also, MOs often have off target effects, and CRISPR approaches are now used to deal with this issue. Due to these issues, I recommend omitting the zebrafish experiments and following this up in a future more complete study. In addition, Figure S3C of β-cardiac myosin and NMIIB staining in heart sections could simply be showing NMIIB in a small gap between two adjacent cardiac myofibrils. One would like to see a linear array of the BCMII/NMIIB along some continuously stained F-actin fibers to be convinced. This could be omitted also.

10) I agree wholeheartedly with the authors that their work on the NMIIA and IIB indicates new refined models are necessary and that we should not get hung up on the terminology. The authors may want to refine their model shown in Figure 11 based on data regarding α-actinin2 localization and assembly, as well as the other additional experiments and clarifications I have suggested.

Specific Points for improving Figures:

1) Figure 1F. I can't really see the transition of the MSFs to myofibrils with sarcomeres in Figure 1F. There are too many other myofibrils crossing it and near it. Also, is this indicated "MSF" really an actin arc? It is practically in the middle of the cell. The wide-field example in Figure 1E is better; we need a confocal example similar to this one.

2) Figure 1H. The kymograph in Figure 1H is strange- the retrograde flow of the MSF is convincing, but the purple line through the "sarcomere" seems to be parallel to the myofibrils, which are orthogonal to the MSFs. The authors need to find another example where the myofibrils with sarcomeres are parallel to the MSFs and to the cell edge – similar to Figure 1E, or to the cell and kymograph in Figure 3E-F.

3) Figure 2. The images of Arp2/3 staining in the control and CK666 cells in Panel C appear to show the edge of a spreading cell; with the actin fibers well-behind the edge (a lamellipodium, presumably). However, the image of the cell in Panel A is not the edge of a spreading cell since the actin fibers are right up at the edge. Where is the Arp2/3 in cells like those in Panel A?

4) Figure 3. It is hard to compare the F-actin distribution in Panel A (SMIFH2 inhibition) with that in Panel C for the FHOD3 siRNA experiment. The F-actin in Panel A could be shown at higher magnification and brighter to match that of Panel C.

5) Figure 4. The NMIIA appears to be enriched right at the edge of the cell, while the NMIIB is more abundant further away from the edge. However, the line scan shown in Figure 4B does not match the images. The image in Panel C where the NMIIA and IIB are co-localized also shows that the IIA is relatively more abundant near the cell edge. The line scan should be replaced with data that better matches the images.

6) Figure 4E. The NMIIA vs NMIIB knockdown is cool! The MYH10 knockdown cell appears to form abundant F-actin donut structures that resemble podosomes. I can see them in Figure 6F also. Are they podosomes? If so, they would contain vinculin. What might this mean? Competition between different F-actin structures? Effects of NMIIB in cell adhesion?

7) Figure 6. Can a high mag field of the NMIIB/β-cardiac myosin at the edge of the cell be shown to illustrate better the authors' point that the filaments are smaller and not lined up? The individual filaments in Panel C show the co-assembly, but not the overall pattern of the β-cardiac myosin compared to the NMIIB. Also, does IIA co-assemble with β-cardiac myosin?

8) Figure 6F. The movie of the β-cardiac myosin in the control cells (Video 6) is nice and clearly shows the thick filament assembly at the very edge of the cell, where the NMIIA is located. Since a major point of this figure is that NMIIB helps coordinate the β-cardiac myosin assembly, I would also like to see a live cell movie of the β-cardiac myosin in the MYH10 knockdown cells.

9) Figure 6F-G. The result that the MYH10 KD reduces F-actin assembly but allows β-cardiac myosin assembly to form thick filaments is interesting. However, a caveat is that it might be difficult to image very thin F-actin fibrils due to the very bright podosomes. Staining for α-actinin2 would address whether there are residual myofibril-like scaffolding structures; as might be suggested by the end-to-end lining up of the β-cardiac myosin structures.

10) Figure 6H. The β cardiac myosin appears to form star like structures in the Latrunculin treated cells. What might these be? If the cells are stained for F-actin, are there residual F-actin fibrils connecting the stars? Where is the α-actinin2? This reminds me of the polygonal arrays of F-actin and α-actinin2 in spreading cardiac myocytes reported by Holtzer's group. Lin et al., 1989. Does Latrunculin lead to changes in cell spreading? Or size?

11) Figure 7. Can the distinction between myosin stack formation by expansion versus concatenation mechanisms be explained more clearly? I don't get it. In the expansion image sequence in Figure 7D, it appears that a thick filament is splitting into two separate adjacent thick filaments- this could later lead to a branched myofibril (a well-known process in myofibril assembly). I don't see how this image shows that the thick filaments are adding on to one another make a thicker myofibril.

Reviewer #2:

This study by Fenix et al., builds on decades-old models of muscle sarcomere formation that postulated that non-muscle myosin assemblies served as templates for the formation of muscle myosin II-containing sarcomeres. This group was optimally positioned to tackle this issue that included important questions, for example the existence of actomyosin filaments containing non-muscle and muscle myosin II elements, the differential role of the major non-muscle myosin II isoforms, etc. This study elegantly answers several of these questions. Specifically, the authors demonstrate that: (1) formins are required for sarcomere formation; not so much the Arp2/3 complex; (2) the non-muscle myosin II isoforms play differential roles as sarcomeric precursors; (3) that this is likely the case in vivo.

Overall, I think this is a terrific study, one that definitively merits publication in eLife. Having stated this, I have some issues with several data sets, which in my opinion require some additions, modifications, and clarifications. However, I am convinced the authors can perform these experiments in a reasonable amount of time.

Specific point:

1) In Figure 1G-H-I, it seems clear that MSF and actin arcs are not the same entities in terms of dynamics, e.g. rates of retrograde flow. However, a major distinctive feature of stress fibers is that they connect focal adhesions, thus they are confined to the lower planes of the cell: coverslip interface. I think it would be a good idea to compare the retrograde flow of these structures using TIRF, not spinning disk as they do.

2) In Figure 2, I am fine with the effect of CK-666 in the localization of the Arp2/3 complex (as measured by anti-p34 staining, I assume; it'd be useful to have this stated in the figure legend). However, I am surprised by the lack of effect on the shape of the leading edge, which is very similar. I would like to see kymographs of the leading edge in control and CK-666-treated cells (similar to Figure 3F, but also showing the extension of the leading edge).

3) My main issue with this study pertains to the interpretation of Figure 4E. While I agree some sarcomeres can be seen in Myh9-depleted hiCM cells, the difference between scramble and Myh9 siRNA is too striking to be described as a mere "disorganization" issue. As such, the metric used in Figure 4G is very misleading. I remain convinced Myh9 plays a role in this process, and this description does not adequately address this. To confirm these data, I would like to see data using p-nitroblebbistatin (so they can do dynamics) and a better way to represent the disorganization caused by Myh9 depletion.

4) Figure 4A, the localization of NMII-A and NMII-B is described as equivalent, yet NMII-B remains "stubbornly" behind NMII-A, which is okay. Although I agree that the existence of co-polymeric forms of the two isoforms reflect the existence of a transitional area, the image in Figure 4A and the quantification shown in Figure 4B are dramatically different and may lead to confusion.

5) I do love the experiments depicted in Figure 5. However, I am confused as to their meaning. In the original paper from the Adelstein group (Tullio et al., 1997), a major issue was the size of the cardiomyocytes, which seems unaffected here. A similar result was seen with the conditional II-B/-C ablation in the Ma et al. paper from 2009. Is the size of the cells significantly larger in these experiments?

6) How is betaCMII organized before sarcomeres form? Does it form mini-filaments? In the same direction, if betaCMII is depleted, does NMII-B distribute evenly throughout the cytoplasm? Or is it still confined to the leading edge?

Reviewer #3:

This paper addresses some very interesting questions regarding sarcomere formation using a nice model cell system. While some of the data looks quite good, a considerable amount has significant issues with quantitation, lack of depth, and/or interpretation. Below are my specific comments.

I am surprised that there is no change in hiCMs after CK666 treatment (Figure 2) other than a reduction in Arp2/3 localization at the cell edge. In most cells Arp2/3 is consuming the bulk of the monomer at steady state. Is there a change in total actin content? Retrograde flow rate? Ability to spread? Do the formin-dependent MSFs get more robust because of the increase in monomer supply when Arp2/3 is inhibited? Of note, their CK666 treatment displaced only ~50% of the Arp2/3 signal from the edge. The remaining Arp2/3 signal/activity could well be enough to drive a significant fraction of normal branched nucleation. I wonder if CK666 works in their cells as expected, and/or if the CK666 in the media after 24 hours of imaging is still active? Does robust KD of any Arp component give the same result? Basically, this aspect of their work needs additional effort before the authors can categorically exclude a contribution from Arp23/ to MSF formation.

I recommend in subsection "Formins but not the Arp2/3 complex are required for MSF-based sarcomere formation…" that the authors move the fourth paragraph in front of the third paragraph. When I read the second paragraph about how SMIFH2 blocks sarcomere formation, I immediately wondered about the MSFs, given that they are supposed to give rise to the sarcomeres. But before I got the answer, I had to read about FHOD3. This seems out of order. In addition, the authors should determine FHOD3's localization relative to MSFs and sarcomeres to see if it supports their idea (and to provide some additional information on FHOD3's role in sarcomere formation since we already know from previous work that sarcomeres disappear when it is knocked down).

The SMIFH2 data in Figure 3 needs a time lapse movie to show how MSFs and sarcomeres disappear, and it needs a washout experiment (preferably with movie) showing the SMIFH2 treatment is readily reversible in these cells.

I think they need at least more incisive experiment to support the idea that MSFs give rise to sarcomeres. One good way to accomplish this would be to use photoactivation or photoconversion to put a fiducial mark in MSFs and then show that the mark persists for some period of time in nascent sarcomeres. This should be readily doable.

The authors use the terminology "MSFs acquire sarcomeres" in various places in the text. To me this does not describe what I think the authors are trying to convey, which is that MSFs transition into sarcomeres or MSFs template sarcomere assembly or something like that. I encourage the authors to think more carefully about their wording.

I am not sure the idea that MIIB is required for sarcomere formation but not for sarcomere maintenance makes sense. I assume that, like everything else, these structures are constantly turning over, so if one blocks sarcomere formation, sarcomeres must disappear over time because of turnover (or the system is more complicated than they think).

I have several significant issues with the data in Figure 4 as regards MIIA KD. First, the authors need to show the level of KD. Without that, one can have confidence in the conclusions. Second, the statement "KD of MIIA did not result in inhibition of MSF or de novo sarcomere assembly" appears to be at the very least a major over-simplification. First, it looks to me like the MSFs are largely missing in the KD cell shown in the middle panel in Figure 4E. Moreover, no quantitation is provided to support the claim that the "KD of MIIA does not result in an inhibition of MSF formation". Second, the effect on sarcomeres looks to be much greater that just disorganization as they call it. Indeed, the scoring for sarcomere assembly in Panel G (percent cells with sarcomeres) is not adequate. Something much more quantitative is required (e.g. total sarcomeres per cell, or per unit). It is my guess that proper quantitation of this data will alter their conclusions significantly.

What I see in Figure 4A and B is very different from what the authors conclude. The authors say that, unlike mesenchymal cells where MIIA is peripheral and MIIB is more central, these two isoforms are in the same place within MSFs of spreading hiCMs. To me, Figure 4A shows very clearly that MIIA is enriched closer to the leading edge than MIIB, exactly as seen in mesenchymal cells. I would need to see simultaneous two-color imaging to be convinced that the distribution MIIA and MIIB in hiCMs is different from that in mesenchymal cells. Similarly, the left most SIM mage in Figure 4C clearly shows (especially on the right side of the cell) mostly red (MIIA) nearest the cell edge, then white moving in (presumably overlap between MIIA and MIIB), and finally mostly green (MIIB) furthest in. My guess is that better analyses and quantitation of all the data in Figure 4A-4C will alter their conclusions significantly. Finally, I suggest the authors determine the ratio of MIIA/MIIB within individual filaments going from the cell edge inward (as in Beach et al., 2014).

I wonder if the MIIB KD effect is the result of not having MIIB or of removing much of the type II myosin in these cells. For example, if MIIB represents 90% of total MII in these cells, then it is possible that the KD effect stems from removing the vast majority of type II myosin, and not specifically MIIB. To address this, the authors need to determine the ratio of MIIA to MIIB (and MIIC?) in these cells. Along these lines, can overexpression of MIIA rescue the MIIB KD phenotype?

I was not clear to me that their quantitation of control and MIIB KD zebrafish hearts is representative because the high mag view of the KD tissue stained with α-actinin in Figure 5B looks pretty much like the high mag view of the control tissue in Figure 5A. In other words, they don't appear to differ by anything like ~3-fold, as presented in the quantitation in Figure 5D. Regarding Figure 5D, I assume that this data was taken from the boxed areas in the images stained for actin in Figure 5C. If I am correct, then this is not appropriate for two reasons. First, the data needs to be collected in a nonbiased way, i.e. not by picking areas but by scoring the entire sample area. Second, because sarcomeres are much clearer in the α-actinin-stained images than in the actin-stained images, the measurements need to be made using α-actinin staining. My guess is that proper scoring of this data will alter the conclusions to some extent.

Regarding Figure 6, the authors say, "We noted that near the leading edge of the cell, BCMII filaments were smaller and not organized into stacks resembling A-bands (Figure 3C)". I do not see data on filament size in Figure 3C. Regarding the conclusion that mixed MIIB/BCMII filaments populate a specific region behind the leading edge, the authors need to provide high mag insets and quantitation of double-stained, SIM imaged cells to support this conclusion. How well was MIIB knocked down in Figure 6F and 6G? I found the conclusion from the latrunculin data- "MSFs and sarcomeric actin filaments are serving as "tracks" with which BCMII filaments are loading onto as they from larger BCMII filament stacks of the A-band"- to be a huge over statement of the data. That things don't work when you blow up all the actin does not even remotely prove this elaborate mechanism. This would require at a minimum very detailed dynamic imaging of the normal maturation process.

Regarding Figure 7, the authors state that they do not "believe MII/BCMII co-filaments are not the major mechanism through which MIIB is facilitating BCMII filament stack assembly". Later, however, they state that "MIIB may be serving as a template to seed the formation of BCMII filaments". I don't see anywhere in the text that the authors clearly articulate what they think the role of MII is in this process. If it is not contributing as a co-filament/template, then what is it doing? If it is contributing as a co-filament/template, then how is their thinking different from the Sanger model? Is there a reason the authors cannot image dynamically MII and BCMII at the same time? It seems to me that the process is slow enough that doing multi-channel imaging would work. Doing this would greatly strengthen this central part of the paper. For example, it would show whether BCMII filaments always appear on MIIB filaments, or if they appear independently? Also, if the authors are looking to combine two previous models, then maintaining those names is more appropriate (I suggest the Sanger-Stitch Model).

In the end I did not get much sense how MSFs transition/mature into sarcomeres. Some additional insight into this would strengthen the paper considerably. For example, where is capping protein in this process? At some point, I would expect CapZ to incorporate to build a functional sarcomere. Can the authors shed some light on this part of the process with some staining and/or dynamic imaging of capping protein?

Please use pagination. Please drop the italics on "after" in the third paragraph of the Discussion section. The paper contains a lot of missing words, repeated words, etc. The Introduction could be made significantly more readable with some effort.

Finally, after the submission of this manuscript, a paper appeared in Dev. Cell from Chris Chen's group addressing this same topic. Notably, they came to very different conclusions regarding a role for NMII. While I think disagreement is very often good for science, I would expect any future versions of this manuscript to contain extensive discussions about these differences (and similarities) and possible explanations for the discrepancies.

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

Thank you for submitting your article "Muscle specific stress fibers give rise to sarcomeres in cardiomyocytes" for consideration by eLife. Your article has been reviewed by Anna Akhmanova as the Senior Editor, a Reviewing Editor, and three reviewers. The following individual involved in review of your submission has agreed to reveal her identity: Velia Fowler (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

The revised manuscript is much improved with many new experiments, and substantial rewriting, so that it now presents a coherent story that is well-grounded in the historical context (presented very nicely in the Introduction), answering long-standing questions about the roles of NMIIA and NMIIA in sarcomere formation, using elegant live cell and super-resolution light microscopy approaches in human induced pluripotent stem cell cardiac myocytes (hiCM). The response to reviewers is very thorough and thoughtful. The authors have now used Lifeact-FPs and α-actinin2-FPs to study the transitions of the 'muscle stress fibers' (MSFs) into the myofibrils as the MSFs move dorsally and used the α-actinin2 spots/Z line distances to quantify sarcomere formation very convincingly. They also use photo-activation of Eos-α-actinin2 to show that the α-actinin2 dots in the in the MSFs become Z lines in the sarcomeres. The work shows that the two non-muscle myosin IIs are both important but play somewhat different roles in sarcomere and myofibril assembly, and also that formin FHOD3 but not Arp2/3 actin nucleation is required for sarcomere assembly. The controls are all in place and the conclusions are strong. The authors also show that NMIIA and NMIIB can form heteropolymers with the β-cardiac myosin II (betaCMII) in the cultured hiCMs, a novel finding. The comparisons of the behavior of the arc-like MSFs in the hiCMs with the actin arcs in spreading non-muscle cells such as U20S cells are also interesting and important to the field. Their careful presentation of the outstanding questions and their own new data in a historical context is valuable and balanced.

Essential revisions:

1) While most concerns have been addressed by the substantial, and high quality new experiments and extensive rewriting, there were still a few concerns with the figure presentation and some of the interpretations. Most importantly, the major messages of the manuscript (MSFs are precursors to sarcomeres, NMII participates in sarcomere formation in a precursor role) became somewhat diluted in the revised version. A few conclusions are also somewhat overstated. Below, we suggest improvements to clarify and strengthen the results and conclusions. These can all be addressed by minor revisions to figures and text.

2) It appears from the data shown in the paper that the new sarcomeres are formed by rearrangement of the MSFs, starting at the spreading edge of the cell. The preexisting sarcomeres present before replating appear to be in the center of the replated cells so that they do not play a role in the formation of the new sarcomeres. Since this is not entirely clear, the authors could provide better 3D imaging of a newly replated cell to show where the preexisting sarcomeres are, and their relationship to the newly forming sarcomeres at the edge of the cell.

3) Based on the photoconversion experiments using α-actinin-2, if the authors were to do similar experiments photoconverting NMIIA and/or IIB, where would it go? This is important because of their observation that NMII-A and IIB do not incorporate into the sarcomere. It would be nice, if possible, to address this point experimentally.

4) The zebrafish data is rather weak, and furthermore does not address the main message that "MSF are precursors to sarcomeres, NMII participates in sarcomere formation in a precursor role", and thus it can better be omitted.

Suggestions for revisions of text and figures:

1) Figure 6, Panel E. Very nice data, but the images would be clearer if the individual channels were shown in gray scale, with a merge for both the low and high mag crops included. Also, the high mag crops on the right would be better presented horizontally (i.e., time along X axis), so that we can follow the assembly over time more easily. The authors should comment on their intriguing observation that the FHOD3 appears to assemble into a pair of widely spaced stripes in the middle of the sarcomere. It is not at the Z line, based on the F-actin staining, nor at the pointed end - which would be one stripe in the middle of the sarcomere in the unstretched sarcomeres of the cultured cardiomyocytes. Is the FHOD3 staining similar to previous studies of FHOD3 localization? Or is it new? It looks as if it is in the region of A-I junction, near the locations of the myosin heads. This could be commented about.

2) Figure 8. Very interesting experiments! The effects of MYH9 knockdown versus MYH10 knockdown are very different. While they both are important for sarcomere/myofibril assembly, the MYH9 KD looks more important for lateral myofibril alignment – the KD cells have many branched and wispy looking myofibrils, although they clearly can make sarcomeres; the quantification shows this too. However, the MYH10 KD cells don't make any sarcomeres at all, so NMIIB is required for all (initial?) aspects of sarcomere assembly, while NMIIA is not. Figure 8—figure supplement 3 and Figure 8—figure supplement 4 time-lapse images show these differences very clearly also. The results and other parts of manuscript should be rewritten somewhat to clarify and emphasize these intriguing differences between the NMIIA and NMIIB which are very interesting. Have the authors looked at MSF retrograde flow in either of the knockdowns? This could be interesting (but not required). It was difficult to find any mention of this.

3) Figure 9. The zebrafish data remains unconvincing to me. One can find many areas of sarcomeres in the low mag images in the top panels, both in the myh9bMO and myh10MO hearts. The high mag images in the lower panels are blurry. Moreover, the myofibrils curve in and out of the XY plane in these cells in the heart, and thus some extend in the Z dimension, making it complex to identify all the sarcomeres due to the Z stretch in the confocal. Also, if the myofibrils are even a tiny bit more contracted, then sarcomeres can be very hard to identify from F-actin staining. α-actinin labeling could help. Finally, even if the images and quantification were reliable (which are not convincing to me), the experiment does not address the central point of the study, which is the role of NMIIA and NMIIB in the transition of MSFs to myofibrils containing sarcomeres. There are no clearly distinguishable MSFs in these zebrafish heart cells. We suggest the authors remove this data and save it for a future more extensive study where they use live cell imaging to study the MSF – sarcomere transitions in vivo- probably need lattice light sheet.

4) Figure 10. Panel H. These odd structures that form in the MYH10 KD cells are intriguing. It appears that rods of betaCMII extend between large donut-like foci of F-actin. Where is the α-actinin2? Is it in the center of the F-actin donut? It was speculated in the previous comments that they might be podosomes, and the response to reviewers provided a clearer explanation of why this is unlikely, based on where they are in the cell and what else they may contain. Some this explanation could be incorporated into the manuscript to make the figure clearer to the reader. We are not asking the authors to do more experiments, but at the least they should describe these images more precisely and then speculate about what the structures might be. Also, the betaCMII rods extending between the F-actin donuts could be individual A bands; are they included in the quantification shown in J? Or are A bands only counted if they are in a linear sequence along and F-actin bundle? Panel H high mag panels need a scale bar.

5) Figure 11. Panel I. The image from the Latrunculin-treated cell is confusing. Where are the cells in this image? What is the large fluorescent half-moon at the bottom of the image? Is this an accumulation of β-CMII around the nucleus or center of cell? Are there A bands in this densely-stained area – it looks like there might be. A counter-stain with a nucleus marker or cytoplasmic marker could be helpful. Another question is what type of betaCMII structure is in the little box? Are these 'stars' related to the donut like F-actin structures in panel H? i.e., would F-actin be in the hole in the middle? Was Panel J quantified from I? If so, then wouldn't it make sense to show both the betaCMII and the F-actin staining for the Latrunculin treated cells?

6) Figure 10. The authors' demonstration of NMIIA or NMIIB co-assembly in bipolar filaments with betaCMII in the cultured hiCMs is convincing. However, the reviewers were still not completely convinced by the claim that they can find these co-assembled filaments in vivo in the mouse heart. The heart tissue sections in Figure S3B and Figure S18 are a low mag field of many myofibrils (not sure why Figure S18 is a separate figure, also no scale bar in Figure S3B). Both show that the NMIIB staining (green) blobs are excluded from the myofibrils and squeezed in between the closely packed myofibrils. Are these supposed to be periodic NMIIB filaments located along an MSF type of structure? Or aggregates? One would like to see a zoom in of the blobs region so we can see the NMIIB and betaCM colocalized (along the putative MSFs?) in the context of the heart tissue, ie, we need an intermediate magnification field within which one can find the filaments! Not just the super-resolution images of individual filaments cropped out, as shown.

7) Figure 9—figure supplement 3A. The betaCMII in the siRNA MYH9-treated cells looks remarkably OK. One can see lots of A bands. However, they appear to be lined up along the length of myofbrils that are very thin so that the betaCMII stacks are not evident. It would be nice to show the F-actin colocalization with the betaCMII staining. This also fits with the data in Figure 8 showing that sarcomere formation in MYH9 KD cells is not as disrupted as in MYH10 KD cells (see point 2 above). Presumably the fourier transform does not reveal the periodicity as well as for the controls in Figure 10G, due to the wispy myofibrils and reduced alignment of the stacks. But a line scan along the length of the myofibrils would likely still reveal periodicity. We are not asking for authors to do new experiments here (unless they want to!), rather just tone down interpretation of MYH9 KD images in Figure 9—figure supplement 3, as related to comments in point 2 above about NMIIA function.

https://doi.org/10.7554/eLife.42144.043

Author response

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

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 this paper, the authors investigated sarcomere assembly in human iPSC- derived cardiomyocytes and provide novel molecular insights into this process, which are potentially of broad interest. However, all three reviewers felt that there is a very significant number of issues that need to be addressed before this paper can be published. In particular, the referees found that some data were incomplete and some controls missing, that important quantifications were lacking, and that certain data needed to be analyzed differently. Furthermore, a number of points had been raised that would require addition of new data, such as evaluation of the spacings and dynamics of α-actinin puncta, two-color imaging of myosin II and β cardiac myosin II, and photoactivation experiments to link muscle stress fibers to sarcomere assembly more convincingly. We return the paper to you, hoping that you will find the detailed comments of the reviewers useful for improving your study. However, we will be prepared to consider a new submission, which would fully address the comments of all three reviewers. In case you decide to resubmit your paper to eLife, it will be sent to the same Reviewing Editor, who will then decide, based on your revisions, whether the paper warrants re-review and whether the same reviewers should be consulted.

Reviewer #1:

In this manuscript Fenix et al., have reported a novel phenomenon in sarcomere assembly in human IPS derived cardiomyocytes (hiCM), which builds on prior knowledge in the field. Using confocal, super-resolution and live cell imaging assays in hiCM, the authors demonstrate that sarcomeres assemble from F-actin stress fibers organized in arcs near the edge of the spreading cells, that they term muscle stress fibers (MSF). These resemble actin arcs in spreading cells from HeLa and U2OS cell lines but have quantitative differences. They showed that retrograde movement of the MSFs results in maturation of sarcomeres (i.e. formation of striated myofibrils) and that the rate of retrograde flow of MSFs is significantly slower than that of F-actin arcs in the non-muscle cell types. These observations of myofibril assembly in the hiCMs are overall similar to a number of previous studies using fluorescent-tagged α-actinin to image the transitions of stress fiber-like structures (SFLSs)/pre-myofibrils into myofibrils in primary cardiac myocytes, but numbers and rates have now been quantified here (note the terminology is also different here).

We thank the reviewer for pointing out a number of strengths of this manuscript and acknowledging some of what we have added to the question of sarcomere assembly. We would like to point out that this is the first study we know of to visualize the first sarcomere/myofibril formation event. As the reviewer points out, there are classic studies with montages showing puncta of fluorescently tagged α-actinin 2 joining pre-existing Z lines in myocytes (Dabiri et al., 1997; McKenna et al., 1986). In the original manuscript, we only cited (Dabiri et al., 1997) on this point but, upon further reflection, believe that (McKenna et al., 1986) paper should be cited also. Per all three reviewers’ suggestions below, we now build upon these classic results using photo-activation of α-actinin 2. In addition, to the best of our knowledge no study has visualized the actin cytoskeleton or myosin II during the transition from non-sarcomeric organization to sarcomeric organization in live cells as we show here. The reviewer is correct that many studies (which we referenced in the manuscript) have localized components of putative precursor structures in fixed cells and tissues, but no one has visualized their transition to sarcomeres in live cells as we do here.

We are using the term “MSFs” for two reasons. We apologize for not making this clear in the original version of our Introduction. First, they were named and acknowledged as “stress fiber like structures” by Howard Holtzer years before they were renamed “pre-myofibrils”. We are aware that a portion of the cardiovascular cell biology field uses pre-myofibrils as if the Pre-myofibril Model has been thoroughly tested (Sanger et al., 2005; Sanger et al., 2017). There are just as many, if not more groups that promote other models, such as the Stitching model, as if it too has been thoroughly tested (Chopra et al., 2018; Rui et al., 2010). As such, we feel that calling these stress fibers pre-myofibrils is premature and may lead to those that do not “believe” in the Pre-Myofibril Model to dismiss our results outright. Thus, we have chosen to revert back to the simple term “stress fiber” and in the case of this particular manuscript “muscle stress fibers” as we need to distinguish between seemingly similar stress fibers in non-muscle cells and muscle cells. We believe future work which is not directly comparing non-muscle and muscle cells should simply call these stress fibers. This affords an unambiguous discussion of the mechanisms of sarcomere assembly, as using terms such as “pre-myofibril”, “I-Z-I bodies”, or “floating A-bands” implies pre-conceived notions of how the system functions (Holtzer et al., 1997; Rui et al., 2010).

Finally, distinguishing between these models is not a trivial point, especially in light of the recent work by (Chopra et al., 2018). This manuscript was published in a top tier journal and failed to properly introduce or discuss any previous models of sarcomere assembly, such as those mentioned above. Indeed, they present what is more or less the Stitching model but with the added twist of sarcomeres streaming from the adhesions. Although, this in itself is reminiscent of the previous proposal from (Quach and Rando, 2006) in skeletal muscle- also not cited by (Chopra et al., 2018). We of course are not addressing the relationship between adhesions and myofibrils in our current manuscript as it is well outside the scope of this study and is an ongoing project in our lab. We merely mention this in the response to emphasize that a lack of discussion of the previous models leads to proposals of “new” models. We feel this simply muddies an already dense and confusing literature. As such, we have revised our Introduction to introduce the Major models in the fields more explicitly and in the Discussion section, put our findings in this context.

In a new advance, they go onto show that the formin FHOD3 rather than Arp2/3 plays pivotal roles in MSF directed myofibril assembly/sarcomere formation (by contrast, Arp2/3 is important for assembly of non-muscle cell F-actin arcs). In addition, they demonstrate novel functions for NMIIA versus NMIIA in myofibril assembly, showing that in the MSF to sarcomere transformation process NMIIB is necessary, while NMIIA appears to play a redundant role, using an acute knockdown approach. Finally, the mechanism of β-cardiac myosin II thick filament assembly into A bands was also studied, and live cell imaging was used to evaluate the mechanism of thick filament assembly into sarcomeres in striated myofibrils. Through these experiments Fenix et al., have convincingly shown that MSFs (which likely correspond to previously-termed SFLSs or pre-myofibrils) are precursors for maturing sarcomeres and require a different non-muscle myosin isoform as compared to the arc-like stress fibers found in non-muscle cell lines.

To clarify, we have never claimed that the MSFs were not corresponding to the previously described structures. We are grateful to the reviewer for pointing out that one could interpret our wording in a way that we are claiming novelty. Our intent here is to build upon the work of others, which we cited in the previous version of the manuscript and are now more explicit in our Introduction and Discussion section so that the reader understands the previous work and our perspective.

However, some data are weak and do not add to the main points (e.g., the zebrafish data), and there are some missing pieces in the data (e.g., lack of direct comparisons to the previous literature using α-actinin2 as a marker for myofibril assembly, questions regarding location of FHOD3, potential myosin isoform compensation), that need to be addressed to place this study in the context of previous work. Many of the figures also need to be improved to strengthen the points and clarify some ambiguous results.

We thank the reviewer for pointing out a number of points that need addressing. Doing so has made our manuscript stronger. We address these points in depth below.

Main points:

1) The authors claim in the Introduction that the hiCM system allows them to study de novo myofibril assembly in vitro for the first time, unlike the previous primary cardiac cell cultures from embryonic chick hearts or neonatal mouse hearts. However, Figure 5 shows that the hiCM cells contain sarcomeres before replating and spreading! Thus, the hiCM system is actually quite similar to the primary cell systems, which disassemble and then reassemble sarcomeres after replating and spreading.

We partially agree with the reviewer on this point and have revised our language about the primary cultures of embryonic cardiomyocytes. We have removed “de novo” from the manuscript. We did not mean to imply that primary cardiomyocytes (e.g., embryonic) do not form sarcomeres de novo after isolation and re-plating. They probably do, and we suspect that several groups have unpublished data showing this point. However, the data in the published literature using these systems to visualize and test the mechanisms of sarcomere assembly has thus far only provided data after cells contain sarcomeres. Here, we utilize a re-plating protocol, which was not trivial to establish, to visualize the first sarcomeres assembled in live cardiac myocytes. Nevertheless, we have softened our language in the current version of this manuscript considerably on this point.

Of note, the recent report by (Chopra et al., 2018) has also used the term “de novo to refer to sarcomere assembly throughout their manuscript. As discussed at the end of this response, we believe there are significant problems with (Chopra et al., 2018) paper and they are, indeed, not showing the first sarcomeres that assemble in hiCM. Nonetheless, we have removed the term “de novo” in lieu of more descriptive wording.

Nevertheless, the hiCM cells are a great system, and the authors present some interesting and novel findings regarding the roles of actin arcs and myosin isoforms in myofibril assembly. The Introduction and Results section should be rewritten and shortened to emphasize the new work on formins and myosin isoforms.

As implied above, we have re-written the Introduction, Results section and Discussion section of the current manuscript to more clearly emphasize our results within the context of the larger literature.

2) There is another very important point regarding previous work on myofibril assembly. The authors state that they are unable to find I-Z-I bodies in the hiCMs, but I can clearly see small α-actinin puncta along the F-actin arcs in Figure 2A. The authors may have been misled by the vast and confusing literature; I-Z-I bodies are simply small α-actinin2 puncta linearly arranged along F-actin bundles that stain continuously for F-actin.

We agree with the reviewer that there is little evidence in the literature as proposed in the Stitching model. Indeed, others seem to find this confusing as well as the localization of α-actinin 2 puncta has been used to discuss both the Pre-myofibril (Kan et al., 2012) and Stitching Models (Rui et al., 2010). Therefore, we have chosen to test the prediction of relative positions of assembling sarcomeres as proposed by the Templating/Pre-Myofibril Models and Stitching Model. There is no discussion of I-Z-I bodies in the current Results section or Discussion section of our manuscript.

These do appear to be present in the hiCMs. The authors should reexamine their control (untreated) hiCMs and present some high mag images of the cell edge and the F-actin arcs stained for α-actinin2. How does the spacing of α-actinin2 puncta in the arcs/MSFs of hiCMs compare with the literature on myofibril assembly in other cardiac cell types? The distances between the α-actinin2 puncta should be measured in the various experiments, including the formin and myosin KD experiments.

We thank the reviewer for this comment as it has substantially strengthened our manuscript. In the original submission of our manuscript, we did not want to exhaustively characterize the α-actinin 2 in MSFs of hiCMs, as this has been thoroughly characterized in other systems as the reviewer mentions. We now provide thorough characterization of α-actinin 2 spacing in hiCMs in control (untreated), and all experimental conditions. This has not only led to a more thorough characterization and quantification of MSFs, but also a more complete understanding of the effects of our perturbations. Most importantly, these additional quantifications will more easily allow others to repeat our experiments and compare their own experimental results to what we present here.

It is important to compare the so-called MSFs in this study with the various so-called "SFLSs" and "pre-myofibril" etc structures studied by other groups. Most of the live imaging in previous work used fluorescent-α-actinin2 to study formation of myofibrils; thus, one would like to know whether this study obtains similar results.

We are confused. Before submission of our original manuscript we conducted a thorough review of the literature. The only live cell examples we could find of live-cell imaging of α-actinin 2 showed cells that already robust Z-lines in the first frame of the montage (Dabiri et al., 1997; McKenna et al., 1986). We also cited one of these papers in the Introduction of the previous version of our manuscript:

“Though a pre-myofibril structure containing actin, NMIIB, α-actinin has never been directly shown to transition into a sarcomere containing myofibril, previous live cell data using fluorescently tagged injected α-actinin has shown “Z-bodies” at the edge come together as the Z-bodies undergo retrograde flow and assemble into existing sarcomeres (Dabiri et al., 1997).”

We believe we have substantially clarified this issue in the current version of the Introduction. Furthermore, we would be grateful to know if we are missing or overlooking literature showing the first sarcomeres assembling in live cells. Of course, there are many images of fixed cells and tissues that were used to predict the Template and Pre-myofibril models.

For example, closely-spaced α-actinin puncta can be seen transitioning into the wider striations in the myofibrils as they move inwards at the spreading edge of a chick cardiac myocyte in Figure 5 in Dabiri et al., 1997. I think these could be the arcs that are shown here in the hiCMs. There are other types of examples in the literature.

We agree with the reviewer that the mentioned example showed nicely that α-actinin 2 puncta transitions into the wider striations of the myofibrils. The reviewer is also correct that in our system the smaller α-actinin 2 puncta do localize to MSFs while the larger α-actinin 2 Z-lines localize to myofibrils. We have now included photo-conversion experiments of α-actinin 2 to repeat and expand on the results of Dabiri et al. (New Figure 2). Again, we never meant to make it sound that MSFs were novel structures.

3) As a related point, what is the direct evidence that the F-actin arcs in the hiCMs are similar to the F-actin arcs in non-muscle cells? The first paragraph of the Results section states: "Strikingly, super-resolution imaging revealed the MSFs in hiCMs resembled a classic actin stress fiber found in non-muscle cells, referred to as actin arcs (Figure 1C and 1D)." I expected the figure would show high mag images of the hiCM arc substructure compared to the non-muscle cell arcs, but this figure shows relatively low mag images of the arcs in U2OS and HeLa cells, revealing continuous F-actin along the fibers in all cell types. How does this show that the MSFs are similar to the arcs in the non-muscle cells? We need to see some high-resolution images of the arcs in the hiCMs and the non-muscle cells--for example what are the spacings between the Z bodies (α-actinin puncta) in the arcs in each cell type? I would like to know whether the NMIIA and NMIIB are located in between the α-actinin2 puncta of the MSFs, which would also be similar to the SFs in the non-muscle cell types (but with a different α-actinin isoform).

We thank you for this comment. There are several similarities between actin arcs and MSFs that we failed to articulate. First, they both exist on the dorsal surface of the spreading cells. They stain along their lengths continuously with phalloidin indicating that there are few if any regions without actin. They are also both arranged parallel to the edge of the cell and move in a retrograde manner away from the edge. We have substantially revised Figure 1 and first part of the Results section to show evidence for these points.

4) Actin arcs are shown in the live cell imaging experiments to be precursors for "myofibrils containing sarcomeres" (note- this is a more precise terminology- a sarcomere is a small unit of a myofibril- and the arc may have a small actomyosin 'mini-sarcomere' type structure similar to the stress fibers of non-muscle cells). Can other types of actin stress fibers like dorsal and ventral stress fibers also mature into sarcomeres in hiCM? Or, do these hiCM cells only possess actin arcs? This seems unlikely, as I think I can see some radial stress fibers in the cells in Figure 3E and in Figure 1H near the edge.

There are several actin filament-based structures in hiCMs that we are not focusing on in this particular manuscript. Our data does indicate that there are similar structures in spreading hiCMs to the three major types of stress fibers in non-muscle cells: actin arcs (sometimes called transverse arcs), dorsal stress fibers and ventral stress fibers. With that said, we have not detected sarcomeres arising from either the dorsal stress fiber-like population or the ventral stress fiber population. It is important to note that our manuscript focuses on the events that occur before the hiCMs have been on the plate for 24 hours. There are several interesting changes that occur post 24 hours. Sarcomeres do appear on the bottom of the cell and sometimes along what appear to be dorsal stress fibers. Ongoing work in our lab is addressing these changes. In the current manuscript we do show two examples of sarcomeres appearing on the ventral surface of the cell and is more thoroughly discussed in response to reviewer 3’s final point.

5) The authors have shown that FHOD3 knockdown in hiCM cells results in reduced sarcomere formation. However, they have not shown the cellular distribution of FHOD3 by immunofluorescence, unlike Arp2/3. Where does FHOD3 localize in the hiCM in the initial MSF stage, and later when the sarcomeres are formed? Does FHOD3 directly interact with the MSFs and/or the sarcomeres?

We thank the reviewer for this point. Yes, FHOD3 is localized to both MSF and sarcomeres (Figure 6E). We now include localization of a FHOD3-mEGFP construct, which was a gift from and previously published by Dr. Elizabeth Ehler’s group.

6) The authors use a pan-formin inhibitor, SMIFH2, to show that formins are important for formation of the actin arcs and sarcomeres. While FHOD3 is clearly important, their experiments do not rule out roles for other formins. For example, see Rosado et al., 2014. Also, the authors claim that mDia1 is not involved in the MSFs, but this has not been tested here. This could be tested, or at the least, discussed and clarified.

We have used RNAseq to identify the formins expressed in the hiCMs used in this study (Figure 6A). As we report in the revised version of the manuscript, there are three highly expressed formins in hiCMs; FHOD3, DAAM1, and mDia1. These are actually different from the most highly expressed Formin paralogs in mouse hearts as reported by Rosado et al. We now provide a discussion of these results in the Discussion of the current manuscript. Furthermore, we have knocked down both DAAM1 and mDia1. Both knockdowns produce clear phenotypes, which warrant future investigations (Figure 6—figure supplement 1).

We did not mean to imply that mDia1 is not involved in MSFs. The only mention of mDia1 was as follows in the Discussion section:

“However, we found that the formin paralog, FHOD3, was responsible for MSF and sarcomere assembly, whereas in non-muscle cells mDia1 is required for actin arcs (Murugesan et al., 2016).”

We do understand how this sentence could appear that we are implying mDia1 is not required for MSFs or sarcomere assembly. That was not our intention and it has been removed.

7) NMIIA and NMIIB can form heteropolymers in hiCM, yet NMIIB knockdown alone disrupts sarcomere formation. Is this because there is a compensatory rise in NMIIB expression upon NMIIA knockdown, which restores the total NMII levels in the cell? However, another possibility is that the NMIIA levels are not enhanced upon NMIIB knockdown, resulting in a drop of cellular total NMII levels, bringing about a disruption of sarcomere assembly. The authors should examine the NMIIA levels upon NMIIB knockdown, and vice versa, to address this issue. It would also be helpful to provide a quantitative estimate of NMIIA/NMIIB distribution in the soluble and insoluble fractions of hiCM cells during the MSF and sarcomere stages. This may confirm why one isoform is more important than the other in sarcomere formation.

We thank the reviewer for these points. Of note, there has been extensive changes to our interpretation of the results from the NMIIA KDs in hiCMs based on comments from all three reviewers. We now provide quantifications which suggest NMIIA is playing major roles during sarcomere assembly in hiCMs (Figure 8). We also provide protein level changes of NMIIA levels upon NMIIB knockdown and vice-versa (Figure 8—figure supplement 2). The reviewer’s comments on further investigating the mechanism of how NMIIA and NMIIB drive sarcomeres formation is a good one. We have begun another project mapping the domains of each myosin that is required. While overexpression of NMIIA does not rescue we have found that we can rescue with only the motor domain of NMIIB (i.e., a chimera with the rod of NMIIA and the motor of NMIIB). Immuno- fluorescence was used to confirm that endogenous NMIIB was knocked down in each expressing cell. While these results are interesting it has become clear to us that the role of each domain and specific levels of each in filaments and the cytosol is beyond the scope of our current study. The reviewer is asking for at least one (maybe more) follow-up studies to be added to an already massive manuscript.

8) Although the results show that NMIIB is essential for de-novo sarcomere assembly in spreading cells, is it also important for the maintenance of the sarcomeres?

No, our data would suggest that neither NMIIB or NMIIA are required for sarcomere maintenance, at least over the course of 11 days in culture. We provided this data for NMIIB in Figure 5A of the previous version of this manuscript. Due to comments from all reviewers it was clear that this was a very confusing way to present this data, as it was included in the same figure as the Zebrafish data. The maintenance data is now included in Figure 8F and Figure 8—figure supplement 6. While our data suggests there may be separate mechanisms of assembly and maintenance, which is not necessarily a surprise, we feel it is outside the scope of this current manuscript to delineate these mechanisms.

9) I appreciate the authors' efforts to study NMIIB in vivo, but the experiments are not convincing. The zebrafish morpholino experiments are potentially exciting but are underdeveloped and missing important controls. What is the result of MO for MYH9 (NMIIA)? MO of the MYH10 (NMIIB) could also affect heart development via effects on non-cardiac cell types, e.g., vascular endothelium. A rescue using a cardiac-specific MYH10 transgene would address this. Also, MOs often have off target effects, and CRISPR approaches are now used to deal with this issue. Due to these issues, I recommend omitting the zebrafish experiments and following this up in a future more complete study.

We disagree the zebrafish data should be omitted. As mentioned above, comments from reviewers 2 and 3, caused us to re-evaluate the role of NMIIA during sarcomere assembly.

This included knocking down NMIIA via MO in zebrafish as reviewer 1 suggests here. NMIIA KD animals failed to form sarcomeres as assessed by Z-lines which can be clearly seen with an actin stain (current manuscript Figure 9). We also provide more thorough quantification of both NMIIA and NMIIB KD hearts. These included number of sarcomeres, Z-line lengths, and the persistence of sarcomeres along myofibrils for all Zebrafish conditions (current manuscript Figure 9). While the suggested rescue experiment using a cardia-specific transgene is a wonderful idea, it is not realistic or standard in the field of in vivo KDs of motor proteins in KD animals. Furthermore, the proper controls for the off-target effects of MOs were conducted, as previously shown by Dr. Jennifer Gutzman and colleagues. We cited this work in the previous version of the manuscript, but clearly did not include enough detail in our Methods, which we now provide. To further address this concern, we also provide quantification of NMIIA/NMIIB KD efficiency and compensation by the other paralog (Figure 9 and Figure S12). We also agree with the reviewer that further studies based on these data should be undertaken. Indeed, multiple studies have reported less than what we report here yet have stated more. We simply state here that NMIIA and NMIIB are required for sarcomere formation in zebrafish. We make no claims this is cell-type specific.

In addition, Figure S3C of β-cardiac myosin and NMIIB staining in heart sections could simply be showing NMIIB in a small gap between two adjacent cardiac myofibrils. One would like to see a linear array of the BCMII/NMIIB along some continuously stained F-actin fibers to be convinced. This could be omitted also.

This data is not showing NMIIB next to βCMII, it is showing NMIIB and βCMII are in the same filament. We provide multiple examples of these structures in hiCMs, mouse tissue, and human tissue. We used the same technique shown by (Beach et al., 2014) to show multiple myosin II species in co-filaments in cells. As this is the first work showing NMII-βCMII co-filaments in both cells and tissue, this data should not be omitted.

10) I agree wholeheartedly with the authors that their work on the NMIIA and IIB indicates new refined models are necessary and that we should not get hung up on the terminology. The authors may want to refine their model shown in Figure 11 based on data regarding α-actinin2 localization and assembly, as well as the other additional experiments and clarifications I have suggested.

We have done so.

Specific Points for improving Figures:

1) Figure 1F. I can't really see the transition of the MSFs to myofibrils with sarcomeres in Figure 1F. There are too many other myofibrils crossing it and near it. Also, is this indicated "MSF" really an actin arc? It is practically in the middle of the cell. The wide-field example in Figure 1E is better; we need a confocal example similar to this one.

Thank you for this comment. We have now added several more examples of the assembly of the initial myofibril (Figure 1E-G). Some MSF travel further than others before they gain a myofibril like organization of actin and the density of stress fibers/myofibrils make it difficult to clearly see the transition. Figure 1G is an example of this problem. As such, we are only using Figure 1G to show sarcomeres appear on the dorsal (top) surface of hiCM.

2) Figure 1H. The kymograph in Figure 1H is strange- the retrograde flow of the MSF is convincing, but the purple line through the "sarcomere" seems to be parallel to the myofibrils, which are orthogonal to the MSFs. The authors need to find another example where the myofibrils with sarcomeres are parallel to the MSFs and to the cell edge – similar to Figure 1E, or to the cell and kymograph in Figure 3E-F.

We agree and have provided a different example with sarcomeres parallel to the edge. Thank you for pointing this out.

3) Figure 2. The images of Arp2/3 staining in the control and CK666 cells in Panel C appear to show the edge of a spreading cell; with the actin fibers well-behind the edge (a lamellipodium, presumably). However, the image of the cell in Panel A is not the edge of a spreading cell since the actin fibers are right up at the edge. Where is the Arp2/3 in cells like those in Panel A?

The cell in Panel A was treated with CK666 for 24 hours. For the reviewer’s benefit, we have cropped this Figure out using the “snipping tool” in Microsoft Windows and stretched the image in FIJI (ImageJ) (Author response image 1). As can be seen in the stretched image, there is actin all the way to the edge where Arp2/3 is localized. We intentionally displayed the actin at a lower level so that the Arp2/3 localization can be seen clearly.

Author response image 1
Figure 4F stretched.
https://doi.org/10.7554/eLife.42144.036

4) Figure 3. It is hard to compare the F-actin distribution in Panel A (SMIFH2 inhibition) with that in Panel C for the FHOD3 siRNA experiment. The F-actin in Panel A could be shown at higher magnification and brighter to match that of Panel C.

The actin and merge of actin and α-actinin 2 in SMIFH2 has been increased in size (current Figure 5A).

5) Figure 4. The NMIIA appears to be enriched right at the edge of the cell, while the NMIIB is more abundant further away from the edge. However, the line scan shown in Figure 4B does not match the images. The image in Panel C where the NMIIA and IIB are co-localized also shows that the IIA is relatively more abundant near the cell edge. The line scan should be replaced with data that better matches the images.

Thank you for this point. All 3 reviewers brought up this issue. The line scan was created from averaging the localization of NMIIA and NMIIB from multiple individual cells (previous Figure 4B, Current Figure 7B). As such, we chose to change the images to reflect the average. The examples shown in the original manuscript (previous Figure 4A) were from two different cells and not directly comparable. We have replaced the original Figure 4A with localizations of NMIIA and NMIIB in the same hiCM (current Figure 7A).

6) Figure 4E. The NMIIA vs NMIIB knockdown is cool! The MYH10 knockdown cell appears to form abundant F-actin donut structures that resemble podosomes. I can see them in Figure 6F also. Are they podosomes? If so, they would contain vinculin. What might this mean? Competition between different F-actin structures? Effects of NMIIB in cell adhesion?

We think these structures are unlikely to be podosomes (at least not the ones attached to the substrate). They are distributed throughout the cell and are not localized at the ventral surface (Author response image 2). However, there is a likely possibility vinculin and other podosome proteins will localize to them. We do detect podosome-like structures on the ventral surface of control hiCM particularly early after plating. We are currently working on addressing the podosomes more thoroughly in an ongoing project in the lab focusing on how substrate adhesions link to myofibrils.

Author response image 2
3D projection of actin- NMIIB KD hiCM.
https://doi.org/10.7554/eLife.42144.037

7) Figure 6. Can a high mag field of the NMIIB/β-cardiac myosin at the edge of the cell be shown to illustrate better the authors' point that the filaments are smaller and not lined up? The individual filaments in Panel C show the co-assembly, but not the overall pattern of the β-cardiac myosin compared to the NMIIB. Also, does IIA co-assemble with β-cardiac myosin?

Thank you for this comment. This was an obvious oversight on our part. We also did not provide such a view for the co-filaments of NMIIA and NMIIB (current Figure 7D- same cell as previous Figure 4C). We mentioned that almost all of the NMII filaments contained both paralogs but, obviously, failed to show a field of them. We have also now provided a high mag field of endogenous β-cardiac myosin and NMIIA at the edge (Figure 9—figure supplement 2). As shown, it is relatively easy to “find” co-filaments in regions less populated by filaments (Figure 9—figure supplement 2B, boxes 1 and 2). However, in more dense regions it is more difficult to discern them with the spatial resolution afforded by SIM (Figure 9—figure supplement 2B, box 3). We feel this is important information for anyone who will work on the potential roles of co-filaments in the future and thank the reviewer again for this comment.

8) Figure 6F. The movie of the β-cardiac myosin in the control cells (Video 6) is nice and clearly shows the thick filament assembly at the very edge of the cell, where the NMIIA is located. Since a major point of this figure is that NMIIB helps coordinate the β-cardiac myosin assembly, I would also like to see a live cell movie of the β-cardiac myosin in the MYH10 knockdown cells.

We do not see how adding a live cell movie of the β cardiac myosin in the MYH10 knockdown cells would add any new insight, as we already show these cells fail to form Abands comparable to control hiCMs (current Figure 10I).

9) Figure 6F-G. The result that the MYH10 KD reduces F-actin assembly but allows β-cardiac myosin assembly to form thick filaments is interesting. However, a caveat is that it might be difficult to image very thin F-actin fibrils due to the very bright podosomes. Staining for α-actinin2 would address whether there are residual myofibril-like scaffolding structures; as might be suggested by the end-to-end lining up of the β-cardiac myosin structures.

Indeed, β cardiac myosin II filaments are associated with remaining actin structures in the NMIIB KD hiCMs. We failed to mention this in the previous version of the manuscript. We now provide a high magnification view of a βCMII filament on residual actin filaments in the NMIIB KD hiCMs (current Figure 10H). We thank the reviewer for catching this oversight. This association was the reason we performed the subsequent Latrunculin experiments to more thoroughly remove the actin cytoskeleton.

10) Figure 6H. The β cardiac myosin appears to form star like structures in the Latrunculin treated cells. What might these be? If the cells are stained for F-actin, are there residual F-actin fibrils connecting the stars? Where is the α-actinin2? This reminds me of the polygonal arrays of F-actin and α-actinin2 in spreading cardiac myocytes reported by Holtzer's group. Lin et al., 1989. Does Latrunculin lead to changes in cell spreading? Or size?

The star-like structures the reviewer mentions, which we now present a blow-up of (current Figure 10I), are similar in structure to the aggregates of cytoskeletal proteins shown in non-muscle cells treated with Latrunculin or Cytochalasin. These aggregates contain small actin filaments that somehow did not get recycled by endogenous mechanisms during treatment

(precise mechanism unknown). This is probably what the βCMII filaments are binding to. Indeed, in non-muscle cells, NMII and other cytoskeletal components, such as some formins localize to these such puncta (Luo et al., 2016; Luo et al., 2013), while α-actinin does not (Luo et al., 2013).

We did not notice any major changes to cell size. As stated in the text, we treated these cells after they had already spread for 18 hours, not during cell spreading.

11) Figure 7. Can the distinction between myosin stack formation by expansion versus concatenation mechanisms be explained more clearly? I don't get it. In the expansion image sequence in Figure 7D, it appears that a thick filament is splitting into two separate adjacent thick filaments- this could later lead to a branched myofibril (a well-known process in myofibril assembly). I don't see how this image shows that the thick filaments are adding on to one another make a thicker myofibril.

Thank you again for asking for this clarification. We think about the concepts of expansion and concatenation quite often and, as a result, did not notice that we failed to fully describe them in manuscript. We have changed the text to differentiate between these two models with the following text in subsection “NMII and FHOD3 are required for organized A-band formation”:

“In non-muscle U2OS cells, A-band-like stacks of NMIIA filaments are often formed through a process called “Expansion” (Fenix et al., 2016). During Expansion, NMIIA filaments that are close to each other (e.g., in a tight bundle) move away from each other in space but remain part of the same ensemble, where they align into a stack like in the A-band. In addition to Expansion, NMIIA filaments also, but more rarely, “Concatenated” (Fenix et al., 2016).

Concatenation was where spatially separated NMIIA filaments moved towards one another to create a stack.”

Indeed, the reviewer points out that previous Figure 7D (current figure 11D) does not show myosin II Concatenation or the creation of a thicker myofibril. This example now Figure 11D, shows myosin II Expansion, or the movement of filaments away from each other. In hiCMs our data suggests Expansion actually results in a smaller and less organized myosin II ensemble, while Concatenation results in a thicker myofibril which is shown in (previous 7C, current Figure 11C). The arrow heads in current Figure 11C denote βCMII ensembles which concatenate with separate βCMII to create a larger βCMII filament stack.

The phenomenon of myofibril splitting is interesting. However, we rarely see myofibril splitting in live hiCMs. Many phenomena occur at later time points than we are studying and splitting is likely to be among them.

Reviewer #2:

This study by Fenix et al., builds on decades-old models of muscle sarcomere formation that postulated that non-muscle myosin assemblies served as templates for the formation of muscle myosin II-containing sarcomeres. This group was optimally positioned to tackle this issue that included important questions, for example the existence of actomyosin filaments containing non-muscle and muscle myosin II elements, the differential role of the major non-muscle myosin II isoforms, etc. This study elegantly answers several of these questions. Specifically, the authors demonstrate that: (1) formins are required for sarcomere formation; not so much the Arp2/3 complex; (2) the non-muscle myosin II isoforms play differential roles as sarcomeric precursors; (3) that this is likely the case in vivo.

Overall, I think this is a terrific study, one that definitively merits publication in eLife. Having stated this, I have some issues with several data sets, which in my opinion require some additions, modifications, and clarifications. However, I am convinced the authors can perform these experiments in a reasonable amount of time.

Specific point:

1) In Figure 1G-H-I, it seems clear that MSF and actin arcs are not the same entities in terms of dynamics, e.g. rates of retrograde flow. However, a major distinctive feature of stress fibers is that they connect focal adhesions, thus they are confined to the lower planes of the cell: coverslip interface. I think it would be a good idea to compare the retrograde flow of these structures using TIRF, not spinning disk as they do.

Thank you for this comment. We agree that TIRF’s signal-to-noise would be ideal for live imaging of the MSF to myofibril/sarcomere transition. Unfortunately, the transition occurs on the dorsal surface of the cells 1-2 microns away from the substrate. Much like actin arcs in many cells types (e.g., U2OS), the MSF in hiCM travel in a retrograde manner along the dorsal surface of the cell. In our original submission, we failed to articulate the 3D position of the MSF and sarcomeres and showed only showed maximum projections. At the time, it did not seem unreasonable to do this as the top of the cell is relatively flat. However, as the reviewer insightfully points out, not defining the position explicitly will lead to several questions on the reader’s part. As such, we have chosen to revise Figure 1 to include 3D renderings of actin filaments in hiCM. We have also included a time montage in Figure 1G showing the actin on the ventral surface of the cell during the MSF to sarcomere transition. This data is particular important in light of new data published (Chopra et al., 2018) claiming that the first sarcomeres in hiCM form on the ventral surface of the cell at much later time points than we have found.

2) In Figure 2, I am fine with the effect of CK-666 in the localization of the Arp2/3 complex (as measured by anti-p34 staining, I assume; it'd be useful to have this stated in the figure legend). However, I am surprised by the lack of effect on the shape of the leading edge, which is very similar. I would like to see kymographs of the leading edge in control and CK-666-treated cells (similar to Figure 3F, but also showing the extension of the leading edge).

The reviewer is indeed pointing out an interesting phenomenon that has also piqued our interest. The lamellipodium is typically smaller in hiCMs than in some non-muscle cells lines. As such, reducing the hiCM’s lamellipodium is often a minor change. However, this is not always the case. We now also present live-cell data showing P16B-mEGFP and Lifeact-mApple before and after acute treatment with CK666. In addition, as per the reviewer’s suggestion, we have also now added dynamic data from cells treated with CK666. Of note, leading edge dynamics are reduced. In addition, retrograde flow of the MSF is not significantly different. After the revision, this data can be found in Figure 4.

3) My main issue with this study pertains to the interpretation of Figure 4E. While I agree some sarcomeres can be seen in Myh9-depleted hiCM cells, the difference between scramble and Myh9 siRNA is too striking to be described as a mere "disorganization" issue. As such, the metric used in Figure 4G is very misleading. I remain convinced Myh9 plays a role in this process, and this description does not adequately address this. To confirm these data, I would like to see data using p-nitroblebbistatin (so they can do dynamics) and a better way to represent the disorganization caused by Myh9 depletion.

We are indebted to the reviewer for bringing this up and pointing out the weakness of the measurements in what are now Figure 8A and 8B. This has actually resulted in a major reanalysis and change in our interpretation. We did not intend to imply that MYH9 siRNA had a small effect on sarcomere assembly- just that the MYH9 siRNA-treated hiCM had sarcomeres. There is clearly a large effect. We have now stated that there were less sarcomeres and have added new quantifications comparing siRNA scrambled and MYH9 siRNA-treated hiCM. While the distances between Z-lines were not dramatically changed (Figure 8D), there was a significant difference in the length of Z-lines (Figure 8E). We feel these quantifications clarify the phenotype and will make it easier for future studies to directly compare their results to ours.

This line of quantification led us to use MO mediated KD of NMIIA in Zebrafish (Figure 9). MO mediated KD of NMIIA in Zebrafish also resulted in a decrease in the number of sarcomeres (Figure 9C), and sarcomere persistence (Figure 9E). Importantly, the loss of sarcomeres in the NMIIA KD hearts were equivalent. As such, our data suggests that both NMIIA and NMIIB play major roles in sarcomere assembly. Again, we thank the reviewer for this point.

We have also now presented data using blebbistatin. Blebbistatin treatment indeed stopped the assembly of MSFs and sarcomeres (Figure S10).

4) Figure 4A, the localization of NMII-A and NMII-B is described as equivalent, yet NMII-B remains "stubbornly" behind NMII-A, which is okay. Although I agree that the existence of co-polymeric forms of the two isoforms reflect the existence of a transitional area, the image in Figure 4A and the quantification shown in Figure 4B are dramatically different and may lead to confusion.

Thank you for suggesting this clarification. We agree that our language and measurements could lead to confusion. We were trying to convey that neither NMIIA or NMIIB extended as far from the edge as we would expect in non-muscle cells such U2OS. Again, our focus on NMII-B has changed due to the new measurements described in point 3 above. As such, we now report the offset at the edge and then simply state the NMII localization does not persist into regions with sarcomeres. We have also replaced the images of two separate cells with a single cell showing endogenous NMIIA and NMIIB localizations (now Figure 7A). This better reflects the localization and measurements averaged over multiple cells (previously Figure 4B and now Figure 7B). While we did place lone NMIIA filaments at the edge of hiCM in our cartoon model, we clearly failed to articulate the difference in localization. To clarify this issue, we now say in the text on Page 6:

“The vast majority of the NMIIA and NMIIB filaments overlapped, except at the very leading edge where NMIIA is localized slightly ahead of NMIIB in hiCMs (Figure 7A and 7B)”

5) I do love the experiments depicted in Figure 5. However, I am confused as to their meaning. In the original paper from the Adelstein group (Tullio et al., 1997), a major issue was the size of the cardiomyocytes, which seems unaffected here. A similar result was seen with the conditional II-B/-C ablation in the Ma et al. paper from 2009. Is the size of the cells significantly larger in these experiments?

We are hesitant to directly compare our experiments with in vivo experiments with mice. With that said, we do not see larger cells in our experimental model. This is likely due to these cells sitting on a 2D surface.

6) How is betaCMII organized before sarcomeres form? Does it form mini-filaments? In the same direction, if betaCMII is depleted, does NMII-B distribute evenly throughout the cytoplasm? Or is it still confined to the leading edge?

This is a very interesting question. βCMII does indeed form short filaments (similar in length to NMII filaments), specifically at the edge of hiCMs. As shown in Figure10E, the length of βCMII filaments are shorter at the edge and increase in length further away from the edge. We also now present another view of β cardiac myosin II with NMIIA where short filaments of β cardiac myosin II are shown more clearly (Figure 9—figure supplement 2).

The second question the reviewer poses has led to some strange results. First, as further discussed at the end of this response, we do not find that reducing β cardiac myosin II has much effect on sarcomere assembly. In addition, we find that NMIIB stays at the edge even when cells are spread in the presence of blebbistatin (Author response image 3). There is noticeably less NMIIB at the edge, but it does not re-localize throughout the cell as we would expect. This result warrants further investigation and is part of an on-going project in the lab investigating what factors are responsible for the localization of NMIIB (and NMIIA).

Author response image 3
hiCM spread in presence of 100µM Blebbistatin; 24 hours after plating.
https://doi.org/10.7554/eLife.42144.038

Reviewer #3:

This paper addresses some very interesting questions regarding sarcomere formation using a nice model cell system. While some of the data looks quite good, a considerable amount has significant issues with quantitation, lack of depth, and/or interpretation. Below are my specific comments.

I am surprised that there is no change in hiCMs after CK666 treatment (Figure 2) other than a reduction in Arp2/3 localization at the cell edge. In most cells Arp2/3 is consuming the bulk of the monomer at steady state. Is there a change in total actin content? Retrograde flow rate? Ability to spread?

We did not detect a change in total actin content. We have now included experiments measuring the rate of retrograde flow of actin in the presence of CK666, and hiCMs show no change in the rate of retrograde flow in response to CK666 (Figure 4D). As can be seen in Figure 4A, hiCMs are spread similar to control hiCMs in the presence of CK666.

Do the formin-dependent MSFs get more robust because of the increase in monomer supply when Arp2/3 is inhibited? Of note, their CK666 treatment displaced only ~50% of the Arp2/3 signal from the edge. The remaining Arp2/3 signal/activity could well be enough to drive a significant fraction of normal branched nucleation. I wonder if CK666 works in their cells as expected, and/or if the CK666 in the media after 24 hours of imaging is still active?

To quantify sarcomere formation in presence of CK666 (now Figure 4A and 4B), hiCMs were not imaged for 24 hours as suggested by the reviewer. These cells were allowed to spread for 24 hours in the presence of CK666 in the dark as noted in the text and methods, then fixed and stained before imaging. To address the reviewer’s comment and provide a more acute experiment testing the effect of CK666, we imaged live hiCMs expressing a subunit of the Arp2/3 complex, P16B-mEGFP. As can be seen in Figure 4H, treatment of hiCMs with CK666 results in a rapid loss of P16B-mEGFP from the leading edge of hiCMs. In addition, the concentration of CK666 used is well above saturating concentration. Furthermore, inhibition of the Arp2/3 complex via CK666 did not affect retrograde flow rates of MSFs (Figure 4D). Finally, it is important to note that we are not claiming that Arp2/3 plays no role in sarcomere assembly. However, whatever role it might play does not require it to be at the edge.

Does robust KD of any Arp component give the same result? Basically, this aspect of their work needs additional effort before the authors can categorically exclude a contribution from Arp23/ to MSF formation.

We agree with the reviewer that further evidence is required to show that we are inhibiting the Arp2/3 complex. As such, we now provide both acute (Figures 4D and 4H) and long term (24 hours) data showing CK666 treatment results in de-localization of the Arp2/3 complex in fixed and live cells (Figures 4F-I). The reviewer is correct in that we are not categorically ruling out a role for the Arp2/3 complex in sarcomere biology. As such, we have also softened the language we use in our interpretation. The conclusion to subsection “Formins, but not the Arp2/3 complex, are required for MSF-based sarcomere formation” now reads:

“Taken together, our data suggests that the Arp2/3 complex does not need to be localized at the leading edge for sarcomeres to be assembled.”

Given that Arp2/3 plays multiple roles in the cell- including major roles in membrane trafficking, we are hesitant to knock down any individual component. Knock down in our system takes multiple days and we have no way of predicting potential ramifications of long-term removal of Arp2/3 activity.

I recommend in subsection "Formins but not the Arp2/3 complex are required for MSF-based sarcomere formation" that the authors move the fourth paragraph in front of the third paragraph. When I read the second paragraph about how SMIFH2 blocks sarcomere formation, I immediately wondered about the MSFs, given that they are supposed to give rise to the sarcomeres. But before I got the answer, I had to read about FHOD3. This seems out of order.

Thank you for pointing this out. We now see that order of this section was awkward. As such, we now present the SMIFH2 data in Figure 5 and then the FHOD3 data in Figure 6.

In addition, the authors should determine FHOD3's localization relative to MSFs and sarcomeres to see if it supports their idea (and to provide some additional information on FHOD3's role in sarcomere formation since we already know from previous work that sarcomeres disappear when it is knocked down).

Thank you for suggesting this addition. We have now provided the localization of FHOD3 in Figure 6E.

The SMIFH2 data in Figure 3 needs a time lapse movie to show how MSFs and sarcomeres disappear, and it needs a washout experiment (preferably with movie) showing the SMIFH2 treatment is readily reversible in these cells.

We agree with that this data would add further evidence that formin inhibition is preventing sarcomere assembly. We now provided live examples of cells spreading out in SMIFH2 and they do not assemble sarcomeres (Figure S3), and a washout experiment where cells subsequently assemble sarcomeres in (Figure 5—figure supplement 2).

I think they need at least more incisive experiment to support the idea that MSFs give rise to sarcomeres. One good way to accomplish this would be to use photoactivation or photoconversion to put a fiducial mark in MSFs and then show that the mark persists for some period of time in nascent sarcomeres. This should be readily doable.

Thank you for this idea. As reviewer 1 suggested, we have now included experiments and analysis of α-actinin to the manuscript in Figure 2. Along these lines, we now present data showing that some puncta of converted α-actinin-2-mEOS2 in MSF do get incorporate into Zlines.

To address the reviewer’s request, we originally tried to use α cardiac actin for the photoactivation experiments as we thought this might be the most direct way of showing the transition. Unfortunately, this line of investigation did not work. We cloned α cardiac actin from a cDNA library we created from hiCMs and fused it to EOS. This construct did express in hiCMs but did not incorporate well into either MSFs or sarcomeres. We also expressed fluorescently tagged fusions of another actin paralog, βactin, that we know incorporate into actin arcs in U2OS cells. β actin showed a similar localization. It is not wholly surprising that actin monomers with a fluorescent protein hanging off them do not get incorporated into formin-generated actin filaments. Others have noted that fluorescent tagged actin monomers do not seem get past formins well. Nonetheless, we were still disappointed.

The authors use the terminology "MSFs acquire sarcomeres" in various places in the text. To me this does not describe what I think the authors are trying to convey, which is that MSFs transition into sarcomeres or MSFs template sarcomere assembly or something like that. I encourage the authors to think more carefully about their wording.

We agree that transition is a better word for what we are trying to convey. As such, we have modified the manuscript. It should be clear that we believe that MSF are a template for sarcomeres. Thank you for this suggestion.

I am not sure the idea that MIIB is required for sarcomere formation but not for sarcomere maintenance makes sense. I assume that, like everything else, these structures are constantly turning over, so if one blocks sarcomere formation, sarcomeres must disappear over time because of turnover (or the system is more complicated than they think).

Every biological system is more complicated than we think. Such is the burden of those of us who have chosen not to be reductionists. All we can do here is report the data and interpret the best we can. First, NMII filaments are absent from sarcomeres. In addition, after 10 days of KD of NMII (either NMIIA or NMIIB), myocytes have similar sarcomere organization as in siRNA treated controls (Figure 8—figure supplement 6). Based on published data of component turnover in mature sarcomeres, this is enough time for the sarcomeres to turn over several times. However, when asked to assemble sarcomeres without pre-existing sarcomeres in our spreading assay, the NMII KD hiCMs fail to do so. Over all, we think this data supports the proposal that there could be separate mechanisms governing sarcomere assembly and sarcomere maintenance. Clearly, quite a bit of future work will be needed to fully address the molecular mechanisms underlying the molecular differences between assembly and maintenance.

I have several significant issues with the data in Figure 4 as regards MIIA KD. First, the authors need to show the level of KD. Without that, one can have confidence in the conclusions. Second, the statement "KD of MIIA did not result in inhibition of MSF or de novo sarcomere assembly" appears to be at the very least a major over-simplification. First, it looks to me like the MSFs are largely missing in the KD cell shown in the middle Panel in 4E. Moreover, no quantitation is provided to support the claim that the "KD of MIIA does not result in an inhibition of MSF formation". Second, the effect on sarcomeres looks to be much greater that just disorganization as they call it. Indeed, the scoring for sarcomere assembly in Panel G (percent cells with sarcomeres) is not adequate. Something much more quantitative is required (e.g. total sarcomeres per cell, or per unit). It is my guess that proper quantitation of this data will alter their conclusions significantly.

Thank you for bringing this up and pointing out the weakness of the measurements in what are now Figure 8A and 8B. This has resulted in a major re-analysis and change in the interpretation of the data. We did not intend to imply that MYH9 siRNA had a small effect on sarcomere assembly- just that the MYH9 siRNA treated hiCM had sarcomeres. There is clearly a large effect. We have now stated that there were less sarcomeres and have added new quantifications comparing siRNA scrambled and MYH9 siRNA treated hiCM. While the distances between Z lines were not dramatically changed (Figure 8D), there was a significant difference in the length of Z-lines (Figure 8E). We feel these quantifications clarify the phenotype and will make it easier for future studies to directly compare their results to ours.

This line of quantification in hiCM also led us to use MO mediated KD of NMIIA in Zebrafish (Figure 9). MO mediated KD of NMIIA in Zebrafish also resulted in a decrease in the number of sarcomeres (Figure 9C), and sarcomere persistence (Figure 9E). Importantly, the loss of sarcomeres in the NMIIA KD hearts were equivalent to that of NMIIB KD. Combined, the hiCM quantifications and the Zebrafish results have led to a major change in our interpretation.

Clearly our data suggests that both NMIIA and NMIIB play major roles in sarcomere assembly. Again, we thank the reviewer for this point.

What I see in Figure 4A and B is very different from what the authors conclude. The authors say that, unlike mesenchymal cells where MIIA is peripheral and MIIB is more central, these two isoforms are in the same place within MSFs of spreading hiCMs. To me, Figure 4A shows very clearly that MIIA is enriched closer to the leading edge than MIIB, exactly as seen in mesenchymal cells. I would need to see simultaneous two-color imaging to be convinced that the distribution MIIA and MIIB in hiCMs is different from that in mesenchymal cells. Similarly, the left most SIM mage in Figure 4C clearly shows (especially on the right side of the cell) mostly red (MIIA) nearest the cell edge, then white moving in (presumably overlap between MIIA and MIIB), and finally mostly green (MIIB) furthest in. My guess is that better analyses and quantitation of all the data in Figure 4A-4C will alter their conclusions significantly.

We apologize for the confusion. Previously, we presented NMIIA and NMIB in different cells (previous Figure 4A). As such, it was not possible to directly compare the relative localizations of NMIIA and NMIIB. This is why presented quantification of their localizations NMIIB averaged over several cells (previous Figure 4B; current Figure 7B). We did not intend to imply that NMIIA was not slightly in front. We even included this detail in our cartoon model. However, we clearly chose to show cells that were not completely representative of the quantification. In particular, the NMIIB localization that was previously shown in Figure 4A was slightly further from the edge than average. To eliminate confusion on this point, we now present NMIIA and NMIIB localized in the same cell (current Figure 7A). This required us to label one of the antibodies—to NMIIA— directly with a fluorophore.

The main point we were trying to make was that NMIIA/NMIIB was not about which paralog showed up first but that neither extended as far as one would see in mesenchymal cells. We have removed this comparison.

Finally, I suggest the authors determine the ratio of MIIA/MIIB within individual filaments going from the cell edge inward (as in Beach et al., 2014).

This is a confusing request. Beach et al., 2014 did not measure the ratio of MIIA/MIIB in individual filaments, nor did Shutova et al., 2014, which also presented a characterization of NMII co-filaments. (Beach et al., 2017) did not measure MIIA/MIIB ratios either. Beach et al., 2014 did measure NMIIA/NMIIB protein levels following over-expression of the other paralog (using Westerns).

Beach et al., 2014 did report that there were fewer co-filaments of NMIIA/NMIIB further away from the edge. While we do not detect this, it could be because the localization of NMIIA/NMIIB filaments in hiCM do not exist as far from the edge as in U2OS cells.

I wonder if the MIIB KD effect is the result of not having MIIB or of removing much of the type II myosin in these cells. For example, if MIIB represents 90% of total MII in these cells, then it is possible that the KD effect stems from removing the vast majority of type II myosin, and not specifically MIIB. To address this, the authors need to determine the ratio of MIIA to MIIB (and MIIC?) in these cells. Along these lines, can overexpression of MIIA rescue the MIIB KD phenotype?

The reviewer brings up an interesting point here, which is the focus of an ongoing project in the Burnette lab. We also wondered if the effect of NMIIB KD was simply the result of it being the more highly expressed paralog? Based on our preliminary data, we feel that the mechanisms are more complicated. We first performed RNAseq on hiCMs and the mRNA ratio was 1.31:1 for MYH9:MYH10. A difference in turnover kinetics could still result in higher NMIIB protein levels. Therefore, we overexpressed NMIIA after NMIIB KD in three independent experiments, which failed to rescue the NMIIB KD phenotype (response Figure 4). These data have led us to explore what domain of NMIIB is important for sarcomere assembly. We reasoned that the motor domain could be a factor as NMIIA and NMIIB have different duty ratios. To test this, we expressed a chimera with the motor domain of NMIIB fused to the rod domain of NMIIA after NMIIB knockdown. We found that this construct resulted in 80.9 +/- 10.5% of hiCMs containing sarcomeres (response Figure 4B); (N=3 independent experiments; Exp 1: 7/10 cells, Exp 2: 10/11 cells, Exp 3: 9/11 cells). We confirmed that endogenous NMIIB was knocked down in each cell that was analyzed by IF using an NMIIB antibody raised to its C-terminus. While these experimental results are intriguing and encouraging, they are clearly only the start of larger project in which we will need to further characterize these conditions.

Author response image 4
Expression of NMIIA and NMIIA/NMIIB(motor) chimera in NMIIB KD hiCM.
https://doi.org/10.7554/eLife.42144.039

I was not clear to me that their quantitation of control and MIIB KD zebrafish hearts is representative because the high mag view of the KD tissue stained with α-actinin in Figure 5B looks pretty much like the high mag view of the control tissue in Figure 5A. In other words, they don't appear to differ by anything like ~3-fold, as presented in the quantitation in Figure 5D. Regarding Figure 5D, I assume that this data was taken from the boxed areas in the images stained for actin in Figure 5C. If I am correct, then this is not appropriate for two reasons. First, the data needs to be collected in a nonbiased way, i.e. not by picking areas but by scoring the entire sample area. Second, because sarcomeres are much clearer in the α-actinin-stained images than in the actin-stained images, the measurements need to be made using α-actinin staining. My guess is that proper scoring of this data will alter the conclusions to some extent.

We thank the reviewer for these points. We agree the way in which we presented the data in what previously was Figure 5 in the original submission was quite confusing. In the original manuscript, Figure 5A and 5B were hiCMs before re-plating, and Figure 5C and D was from Zebrafish. Though this was noted in the text, figure and figure legend, we agree it was confusing. All data from Zebrafish is now in one main figure (Figure 9) and one supplementary figure (Figure S12). Similar to the NMIIA KD in hiCMs, we have more thoroughly quantified sarcomere assembly from the Zebrafish experiments (Figure 9). As noted above, we also include a NMIIA KD via MO in Zebrafish. In addition, we measure NMIIA and NMIIB protein levels in the KD condition of the other paralog, number of sarcomeres, lengths of Z-lines, and sarcomere persistence (Figure 9C-9E and Figure S12). Importantly, measurements in the original manuscript (previously Figure 5) and the current manuscript (new Figure 9) were not made from the yellow boxed area, which was shown simply to indicate where the high mag image was taken from. As the reviewer notes, it is important to quantify the whole image, which is exactly how these images were quantified. We thank the reviewer for these points, as the Zebrafish data is now much clearer and more thoroughly characterized.

Regarding Figure 6, the authors say, "We noted that near the leading edge of the cell, BCMII filaments were smaller and not organized into stacks resembling A-bands (Figure 3C)". I do not see data on filament size in Figure 3C. Regarding the conclusion that mixed MIIB/BCMII filaments populate a specific region behind the leading edge, the authors need to provide high mag insets and quantitation of double-stained, SIM imaged cells to support this conclusion.

The data was in Figure 6 and Figure S3 and was appropriately cited in the previous version of this manuscript. We presented multiple quantifications to support this statement. First, we presented the localizations of NMIIB and ββCMII in hiCMs and quantified their relative localizations over multiple cells and multiple experiments (previously Figure 6A and 6B). We also measured the distributions of the filament lengths of both NMII and βCMII through the cell (previous Figure 6D and current Figure 10D). We also measure βCMII filament lengths with respect to their position in the cell (i.e., measured from the leading edge of the hiCMs) (previously Figure 6E and current Figure 10E). These measurements suggest that the smaller βCMII filaments are found closer to the leading edge of the cell and largely restricted from the longer βCMII filaments of the A-bands (current Figure 10E). In addition, we do provide hi-mag images of the shorter βCMII filaments (previously Figure 6C and current Figure 10C; previously Figure S3 and current Figure 9- figure supplement 1 and Figure 9- figure supplement 2). These quantifications were all performed from SIM images. All figure legends now explicitly state which imaging modality was used.

How well was MIIB knocked down in Figure 6F and 6G?

On average, we achieved a 70% KD of NMIIB. The quantification of the knock down percentage of both NMIIA and NMIIB is now presented in current Figure 8—figure supplement 2.

I found the conclusion from the latrunculin data- "MSFs and sarcomeric actin filaments are serving as "tracks" with which BCMII filaments are loading onto as they from larger BCMII filament stacks of the A-band"- to be a huge over statement of the data. That things don't work when you blow up all the actin does not even remotely prove this elaborate mechanism. This would require at a minimum very detailed dynamic imaging of the normal maturation process.

We have softened the language simply to, “These results argue that actin filaments are required for the organization of the βCMII filaments of the A-band”.”

Regarding Figure 7, the authors state that they do not "believe MII/BCMII co-filaments are not the major mechanism through which MIIB is facilitating BCMII filament stack assembly". Later, however, they state that "MIIB may be serving as a template to seed the formation of BCMII filaments". I don't see anywhere in the text that the authors clearly articulate what they think the role of MII is in this process. If it is not contributing as a co-filament/template, then what is it doing? If it is contributing as a co-filament/template, then how is their thinking different from the Sanger model? Is there a reason the authors cannot image dynamically MII and BCMII at the same time? It seems to me that the process is slow enough that doing multi-channel imaging would work. Doing this would greatly strengthen this central part of the paper. For example, it would show whether BCMII filaments always appear on MIIB filaments, or if they appear independently? Also, if the authors are looking to combine two previous models, then maintaining those names is more appropriate (I suggest the Sanger-Stitch Model).

We thank the reviewer for catching this statement, as this was a relic of a previous version of the manuscript written before we performed siRNA-mediated knockdown of NMIIA and FHOD3 and localized βCMII filaments (current Figure 9—figure supplement 4). Indeed, we do believe NMII-βCMII cofilaments is a potential mechanism that could lead to A-band assembly and as such this speculation has been removed from the text.

In the end I did not get much sense how MSFs transition/mature into sarcomeres. Some additional insight into this would strengthen the paper considerably. For example, where is capping protein in this process? At some point, I would expect CapZ to incorporate to build a functional sarcomere. Can the authors shed some light on this part of the process with some staining and/or dynamic imaging of capping protein?

No. While we are also fascinated with how MSFs transition/mature into sarcomeres, we strongly feel that pursuing the roles of other proteins in the process is beyond the scope of this manuscript. Future work from our lab and others will investigate interesting potential players such as CapZ. Indeed, even in more rigorously studied systems, such as crawling cells, if/how/when exactly different stress fiber populations transition to one another is not completely understood.

Please use pagination. Please drop the italics on "after" in the third paragraph of the Discussion section. The paper contains a lot of missing words, repeated words, etc. The Introduction could be made significantly more readable with some effort.

We have dropped the italics on “after” in the Discussion section. In addition, a complete rework of the introduction and discussion has been made.

Finally, after the submission of this manuscript, a paper appeared in Dev. Cell from Chris Chen's group addressing this same topic. Notably, they came to very different conclusions regarding a role for NMII. While I think disagreement is very often good for science, I would expect any future versions of this manuscript to contain extensive discussions about these differences (and similarities) and possible explanations for the discrepancies.

We agree with the reviewer that we should address the paper from Chris Chen’s group in our manuscript. As such, we have provided the following in the Discussion section of the current manuscript including two examples of what we believe they are imaging when they claim that sarcomeres assemble from adhesions. Please see below for additional comments on this manuscript we have provided for the Editor and reviewers’ consideration.

Text from the Discussion section.

“Our data shows the transition of a MSF to a sarcomere-containing myofibril occurs on the dorsal (top) surface of the cell. However, a recent study also imaging iPSC derived cardiac myocytes claims that sarcomeres are formed on the ventral (bottom) surface of the cell near extracellular matrix adhesions (Chopra et al., 2018). This group also report that the first sarcomeres forms between 24-48 hours after plating, well after we detect the first sarcomere appearing (Chopra et al., 2018). This led us to question what could be leading to these two seemingly opposite results. Importantly, it appears that this group was imaged the ventral (bottom) surface of their myocytes. Close inspection of their time-lapse movies revealed faint structures corresponding to sarcomeric pattern that show up in the frame right before the appearance of sarcomeres. This supports the notion that they are imaging sarcomeres that are coming into focus, and not assembling “de novo”. To test this idea, we imaged hiCMs with 3D confocal microscopy after they had been plated for 24 hrs (Figure S17). While we also see similar patterns of sarcomeres appearing on the ventral surface as (Chopra et al., 2018), our data revealed these sarcomeres are moving down from the dorsal surface and not assembling on the ventral surface (Figure S17 and Supplemental Movie 3). Of interest, the phenomenon of actin arcs moving down to the ventral surface of non-muscle cells has also been reported previously (Gao et al., 2012; Hotulainen and Lappalainen, 2006). Finally, (Chopra et al., 2018) also claim that neither NMIIA nor NMIIB are required for sarcomere assembly. We also find this strange. While their double NMIIA/NMIIB knockout (KO) cardiomyocyte cell line has α-actinin 2 positive structures, they do not contain continuous labeled Z-lines aligned parallel to each other comparable to the control cell line. The authors did not measure Z-line lengths, spacing, or other criteria needed to define sarcomeres. These discrepancies between our study and theirs, including the role of NMII, need to be harmonized, as it will directly affect our interpretation of future in vivo data concerning sarcomere assembly.”

We feel confident in the arguments that we make in the Discussion section of the main text and feel that they clarify the differences between the study by Chen’s group and the study we present here. In our arguments, we have taken their data at face value. However, there are some concerning aspects of their data and analysis that we do not understand. We have chosen not to comment on them in our manuscript, and instead plan to have a dialogue with the corresponding authors directly. With that said, we believe the points below are germane to the current discussion between ourselves and the reviewers and hopefully will give the reviewers an idea of what we intend to do in the future to further address the findings in (Chopra et al., 2018).

To test whether NMIIA and/or NMIIB were required for sarcomere assembly, (Chopra et al., 2018) used NMIIA, NMIIB, and double NMIIA/IIB knockout (KO) cell lines. These cell lines were presented as, “stable CRIPSR knockout cell lines” in their methods. This is curious for a number of reasons. First, as opposed to the TTNtv, MYH6, and MYH7 KO cell lines which were generated in the iPSC cell stage, (Chopra et al., 2018) created the MYH9 and MYH10 (NMIIA and NMIIB, respectively) KO cell lines by performing CRISPR on differentiated cardiomyocytes. Despite a 3-4 day selection with puromyocin, every micrograph presented of these stable KO cell lines contains cells that are clearly positive for NMIIA and/or NMIIB. In fact, 66.7% of the cells shown have robust NMII localization. This is not in line with their Western Blots, which show robust KD of both NMIIA and NMIIB—though no quantification is provided. Secondly, (Chopra et al., 2018) also measured a surprisingly low number of cells for each experimental condition (i.e., 6 cells over 3 experiments for the NMIIA/NMIIB double knock out cell line). We do not know the reason for this but suspect it is because few cells had lower NMII levels. Finally, it is difficult to even access whether the cells (Chopra et al., 2018) highlight are in fact devoid of protein. The images are displayed in such a way that the background is flat black. With that said, we still think that there could be protein localization in these cells as revealed by stretching the images.

(Chopra et al., 2018) claim that neither NMIIA or NMIIB are required for sarcomere assembly. We do not understand how this they arrived at this conclusion. In the NMIIA/IIB double KO, there are clearly no Z lines comparable to control cells or striated myofibrils containing multiple sarcomeres as seen in control cells (Figure 3A and 3B in Chopra et al., 2018). The authors show a blow-up where there are clear diffraction limited α-actinin 2 puncta but no continuous Z line staining. Indeed, the image used to suggest the NMIIA/IIB double KO cardiomyocytes form sarcomeres appears quite similar to the image in the same figure showing that β MHC is required for sarcomere assembly. The only clear difference is that there is less α-actinin 2 signal between the conditions, which is a major component of the sarcomere content analysis Chopra et al. use to quantify sarcomere assembly. This prompted us to more fully investigate their sarcomere content analysis.

While their methods section is brief, Chopra et al. refer readers to a 2015 paper, (Hinson et al., 2015), where the sarcomere content analysis was developed. It is impossible to fully use this analysis, as multiple variables in the equations used are not defined, and strange terms such as “the energy of sarcomere assembly” are provided. The custom ImageJ and MATLAB codes are also not provided. It does not help that the only paper they cite in their methodology for performing a Fourier transform analysis does not actually include a single reference to Fourier transformation in the entire manuscript. We were already in the process of automating the manual measurements we are presenting in our current manuscript. As such, we have decided to pursue our interest in repeating the “sarcomere content” analysis in the context of this ongoing project. For this we will contact the corresponding authors and request their code so that we can first recapitulate their quantifications and compare it to the physical measurements of Z bodies and Z lines that the software we are developing depends on.

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

Summary:

The revised manuscript is much improved with many new experiments, and substantial rewriting, so that it now presents a coherent story that is well-grounded in the historical context (presented very nicely in the Introduction), answering long-standing questions about the roles of NMIIA and NMIIA in sarcomere formation, using elegant live cell and super-resolution light microscopy approaches in human induced pluripotent stem cell cardiac myocytes (hiCM). The response to reviewers is very thorough and thoughtful. The authors have now used Lifeact-FPs and α-actinin2-FPs to study the transitions of the 'muscle stress fibers' (MSFs) into the myofibrils as the MSFs move dorsally and used the α-actinin2 spots/Z line distances to quantify sarcomere formation very convincingly. They also use photo-activation of Eos-α-actinin2 to show that the α-actinin2 dots in the in the MSFs become Z lines in the sarcomeres. The work shows that the two non-muscle myosin IIs are both important but play somewhat different roles in sarcomere and myofibril assembly, and also that formin FHOD3 but not Arp2/3 actin nucleation is required for sarcomere assembly. The controls are all in place and the conclusions are strong. The authors also show that NMIIA and NMIIB can form heteropolymers with the β-cardiac myosin II (betaCMII) in the cultured hiCMs, a novel finding. The comparisons of the behavior of the arc-like MSFs in the hiCMs with the actin arcs in spreading non-muscle cells such as U20S cells are also interesting and important to the field. Their careful presentation of the outstanding questions and their own new data in a historical context is valuable and balanced.

Essential revisions:

1) While most concerns have been addressed by the substantial, and high quality new experiments and extensive rewriting, there were still a few concerns with the figure presentation and some of the interpretations. Most importantly, the major messages of the manuscript (MSFs are precursors to sarcomeres, NMII participates in sarcomere formation in a precursor role) became somewhat diluted in the revised version. A few conclusions are also somewhat overstated. Below, we suggest improvements to clarify and strengthen the results and conclusions. These can all be addressed by minor revisions to figures and text.

2) It appears from the data shown in the paper that the new sarcomeres are formed by rearrangement of the MSFs, starting at the spreading edge of the cell. The preexisting sarcomeres present before replating appear to be in the center of the replated cells so that they do not play a role in the formation of the new sarcomeres. Since this is not entirely clear, the authors could provide better 3D imaging of a newly replated cell to show where the preexisting sarcomeres are, and their relationship to the newly forming sarcomeres at the edge of the cell.

We believe the organization of Figure 1—figure supplement 1 is leading to confusion. We positioned 1.5 hours after plating and 24 hours after plating next to each other for each sarcomere marker we localized. We have now combined the 1.5-hour time points together in Figure 1—figure supplement 1Aand the 24-hour time points in Figure 1—figure supplement 1B. As shown in Figure 1—figure supplement 1A, there are no discernable sarcomeres at the 1.5-hour time points.

3) Based on the photoconversion experiments using α-actinin-2, if the authors were to do similar experiments photoconverting NMIIA and/or IIB, where would it go? This is important because of their observation that NMII-A and IIB do not incorporate into the sarcomere. It would be nice, if possible, to address this point experimentally.

We thank the reviewers for this suggestion. We did photo-convert myosin IIA-mEOS2 and myosin IIB-mEOS2 at the edge of hiCM. Unfortunately, both paralogs of NMII turned over much faster than α-actinin 2, so it was difficult to follow the marked population for long (Author response image 5). We can say that we did not see photo-converted NMIIs move the middle of the cell where sarcomeres typically assemble. As this result was less than satisfying, we tried an alternative approach. We now present two supplemental movies (Video 2 and Video 3) that show the position of NMIIA with sarcomeres assembling visualized with lifeact and the position of NMIIB with sarcomeres assembling visualized with α-actinin 2. While these are not experiments per se, the movies do demonstrate that region where sarcomeres are appearing are largely lacking NMII.

Author response image 5
Time montage of Myosin IIB-/mEOS2 in a hiCM.
https://doi.org/10.7554/eLife.42144.040

4) The zebrafish data is rather weak, and furthermore does not address the main message that "MSF are precursors to sarcomeres, NMII participates in sarcomere formation in a precursor role", and thus it can better be omitted.

We have removed the zebrafish data.

Suggestions for revisions of text and figures:

1) Figure 6, Panel E. Very nice data, but the images would be clearer if the individual channels were shown in gray scale, with a merge for both the low and high mag crops included. Also, the high mag crops on the right would be better presented horizontally (ie, time along X axis), so that we can follow the assembly over time more easily. The authors should comment on their intriguing observation that the FHOD3 appears to assemble into a pair of widely spaced stripes in the middle of the sarcomere. It is not at the Z line, based on the F-actin staining, nor at the pointed end-- which would be one stripe in the middle of the sarcomere in the unstretched sarcomeres of the cultured cardiomyocytes. Is the FHOD3 staining similar to previous studies of FHOD3 localization? Or is it new? It looks as if it is in the region of A-I junction, near the locations of the myosin heads. This could be commented about.

Thank you for these suggestions. We now present the data as suggested. The reviewers are correct in noticing that FHOD3 does not localize to the Z-line. It localizes in two distinct stripes, one on either side of each Z line. To our knowledge, this not been shown before. This is most likely due to the previous used of diffraction-limited imaging. We have now addressed this issue with the following text added to the Discussion section:

“Future work will be required to elucidate the precise mechanism of how and where FHOD3 potentially nucleates actin filaments in sarcomeres. Canonical sarcomeric actin filaments have their barbed ends embedded within a Z-line (see schematic in Figure 1A). As such, this is where we would have predicted FHOD3 would localize. However, our SIM data shows that FHOD3 does not localize to Z lines (Figure 6F, boxes 4, 5 and 6). Instead, FHOD3 appears to localize on either side of each Z-line. This could indicate that FHOD3 is only transiently associated with the barbed ends of canonical sarcomeric actin filaments. Alternatively, there could barbed ends of actin filaments in this region, which could indicate that the organization and/or dynamics of actin filaments within a sarcomere is more complex.”

2) Figure 8. Very interesting experiments! The effects of MYH9 knockdown versus MYH10 knockdown are very different. While they both are important for sarcomere/myofibril assembly, the MYH9 KD looks more important for lateral myofibril alignment – the KD cells have many branched and wispy looking myofibrils, although they clearly can make sarcomeres; the quantification shows this too. However, the MYH10 KD cells don't make any sarcomeres at all, so NMIIB is required for all (initial?) aspects of sarcomere assembly, while NMIIA is not. Figure 8—figure supplement 3 and Figure 8—figure supplement 4 time-lapse images show these differences very clearly also. The results and other parts of manuscript should be rewritten somewhat to clarify and emphasize these intriguing differences between the NMIIA and NMIIB which are very interesting. Have the authors looked at MSF retrograde flow in either of the knockdowns? This could be interesting (but not required). It was difficult to find any mention of this.

Thank you for these suggestions. We have added the following text to the Discussion section to address these differences more clearly. We hope the reviewers do not mind us taking the term “wispy”. It works well as a descriptor. Thank you again.

“We also show that NMIIA and NMIIB are required for proper sarcomere assembly in our hiCM model system. Interestingly, our data suggest that each motor may be playing a different role(s) in sarcomere assembly. NMIIB KD resulted in no detectable sarcomere assembly (Figure 8, Figure 8—figure supplement 4), suggesting that NMIIB could be required for the initial and possibly subsequent steps of assembly. On the other hand, NMIIA KD-hiCMs were able to assemble sarcomeres, although these were wispy with significantly shorter Z-lines (Figure 8 and Figure 8—figure supplement 3). This NMIIA KDphenotype could be a result of several possibilities including a role of NMIIA in sarcomere maturation, alignment, or stability. Obviously, this is not an exhaustive list of potential mechanisms.”

We have not quantified retrograde flow in these conditions but are part of consortium at Vanderbilt that is currently assembling a lattice light sheet. With this low-dose and high-resolution modality, we plan on performing a detailed 3D study of retrograde flow during sarcomere assembly.

3) Figure 9. The zebrafish data remains unconvincing to me. One can find many areas of sarcomeres in the low mag images in the top panels, both in the myh9bMO and myh10MO hearts. The high mag images in the lower panels are blurry. Moreover, the myofibrils curve in and out of the XY plane in these cells in the heart, and thus some extend in the Z dimension, making it complex to identify all the sarcomeres due to the Z stretch in the confocal. Also, if the myofibrils are even a tiny bit more contracted, then sarcomeres can be very hard to identify from F-actin staining. α-actinin labeling could help. Finally, even if the images and quantification were reliable (which are not convincing to me), the experiment does not address the central point of the study, which is the role of NMIIA and NMIIB in the transition of MSFs to myofibrils containing sarcomeres. There are no clearly distinguishable MSFs in these zebrafish heart cells. We suggest the authors remove this data and save it for a future more extensive study where they use live cell imaging to study the MSF – sarcomere transitions in vivo- probably need lattice light sheet.

We have removed the zebrafish data from the manuscript.

4) Figure 10. Panel H. These odd structures that form in the MYH10 KD cells are intriguing. It appears that rods of betaCMII extend between large donut-like foci of F-actin. Where is the α-actinin2? Is it in the center of the F-actin donut? It was speculated in the previous comments that they might be podosomes, and the response to reviewers provided a clearer explanation of why this is unlikely, based on where they are in the cell and what else they may contain. Some this explanation could be incorporated into the manuscript to make the figure clearer to the reader. We are not asking the authors to do more experiments, but at the least they should describe these images more precisely and then speculate about what the structures might be. Also, the betaCMII rods extending between the F-actin donuts could be individual A bands; are they included in the quantification shown in J? Or are A bands only counted if they are in a linear sequence along and F-actin bundle? Panel H high mag panels need a scale bar.

Thank you for these comments. In the methods section, we have now more clearly defined how we quantified how we quantified the betaCMII data. The following text has been added to the Materials and methods section:

“β cardiac myosin II (βCMII) filament and stack quantification

A similar methodology was used to quantify βCMII A-band filament stacks using endogenous βCMII staining and SIM instead of the actin cytoskeleton. A βCMII filament was quantified as the minimum SIM resolvable βCMII unit which had a bipolar organization (a filament with motor domains on each side), as represented in Figure 10A. hiCMs were quantified as containing βCMII A-band filament stacks if they contained even one βCMII filament stack in the cell. A βCMII filament stack was defined as being thicker than the minimum SIM resolvable βCMII filament (Figure 11A), indicating multiple SIM resolvable βCMII filaments. Indeed, by this metric, βCMII filament stacks have more resolvable “motor-domains” than βCMII filaments.”

5) Figure 11. Panel I. The image from the Latrunculin-treated cell is confusing. Where are the cells in this image? What is the large fluorescent half-moon at the bottom of the image? Is this an accumulation of β-CMII around the nucleus or center of cell? Are there A bands in this densely-stained area – it looks like there might be. A counter-stain with a nucleus marker or cytoplasmic marker could be helpful. Another question is what type of betaCMII structure is in the little box? Are these 'stars' related to the donut like F-actin structures in panel H? i.e., would F-actin be in the hole in the middle? Was Panel J quantified from I? If so, then wouldn't it make sense to show both the betaCMII and the F-actin staining for the Latrunculin treated cells?

We have added several things to clarify this data. First, we have denoted the cell body in the latruculin treated cell shown in Figure 9J with an arrow. Indeed, there is clumping of actin and βCMII that occurs near the nucleus. Both networks are dense and chaotic and were not used for the quantification shown in Figure 9K. This is now stated in the figure legend. We have also added a new supplemental figure where we have stained actin, βCMII and DNA in a cell treated with latruculin as the reviewers suggest (Figure 9—figure supplement 3). We agree that there could be A-bands in the cell body. However, it would take higher resolution imaging than SIM to fully explore this possibility. Given that the experiment was simply performed to further test if disrupting actin filament organization also disrupted βCMII filament organization, we feel that further characterization of the clumping in the cell body is beyond the scope of this study.

6) Figure 10. The authors' demonstration of NMIIA or NMIIB co-assembly in bipolar filaments with betaCMII in the cultured hiCMs is convincing. However, the reviewers were still not completely convinced by the claim that they can find these co-assembled filaments in vivo in the mouse heart. The heart tissue sections in Figure S3B and Figure S18 are a low mag field of many myofibrils (not sure why Figure S18 is a separate figure, also no scale bar in Figure S3B). Both show that the NMIIB staining (green) blobs are excluded from the myofibrils and squeezed in between the closely packed myofibrils. Are these supposed to be periodic NMIIB filaments located along an MSF type of structure? Or aggregates? One would like to see a zoom in of the blobs region so we can see the NMIIB and betaCM colocalized (along the putative MSFs?) in the context of the heart tissue, ie, we need an intermediate magnification field within which one can find the filaments! Not just the super-resolution images of individual filaments cropped out, as shown.

Thank you for these comments. We have now added several arrows in, what is now Figure 12B, showing NMIIB localizations that are close to βCMII. As this is a confocal image, we have now added an intermediate SIM image of a human septal muscle along with 4 high mag insets (Figure 12C). A scale bar has been added to Figure S12B.

7) Figure 9—figure supplement 3A. The betaCMII in the siRNA MYH9-treated cells looks remarkably OK. One can see lots of A bands. However, they appear to be lined up along the length of myofbrils that are very thin so that the betaCMII stacks are not evident. It would be nice to show the F-actin colocalization with the betaCMII staining. This also fits with the data in Figure 8 showing that sarcomere formation in MYH9 KD cells is not as disrupted as in MYH10 KD cells (see point 2 above). Presumably the fourier transform does not reveal the periodicity as well as for the controls in Figure 10G, due to the wispy myofibrils and reduced alignment of the stacks. But a line scan along the length of the myofibrils would likely still reveal periodicity. We are not asking for authors to do new experiments here (unless they want to!), rather just tone down interpretation of MYH9 KD images in Figure 9—figure supplement 3, as related to comments in point 2 above about NMIIA function.

Thank you for this comment. Indeed, there are organized A-bands in MYH9 KD condition. We have changed the title of the Results section to: “NMIIB and FHOD3 are required for organized A-band formation”

And changed the text in the Results section to:

“As FHOD3 KD also resulted in disorganized actin filament architecture, we localized βCMII in this condition. Indeed, FHOD3 KD hiCM had disorganized βCMII filaments compared to control hiCMs (Figure 9—figure supplement 4).”

https://doi.org/10.7554/eLife.42144.044

Article and author information

Author details

  1. Aidan M Fenix

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
  2. Abigail C Neininger

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Formal analysis, Investigation, Visualization, Writing—review and editing
    Competing interests
    No competing interests declared
  3. Nilay Taneja

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Formal analysis, Investigation, Writing—review and editing
    Competing interests
    No competing interests declared
  4. Karren Hyde

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Resources, Methodology
    Competing interests
    No competing interests declared
  5. Mike R Visetsouk

    Department of Biological Sciences, Cell and Molecular Biology, University of Wisconsin Milwaukee, Milwaukee, United States
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  6. Ryan J Garde

    Department of Biological Sciences, Cell and Molecular Biology, University of Wisconsin Milwaukee, Milwaukee, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  7. Baohong Liu

    Department of Biomedical Informatics, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
  8. Benjamin R Nixon

    Department of Medicine, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1840-0179
  9. Annabelle E Manalo

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  10. Jason R Becker

    Department of Medicine, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2107-8179
  11. Scott W Crawley

    Department of Biological Sciences, The University of Toledo, Toledo, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  12. David M Bader

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  13. Matthew J Tyska

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Resources, Writing—review and editing
    Competing interests
    No competing interests declared
  14. Qi Liu

    Department of Biomedical Informatics, Vanderbilt University Medical Center, Nashville, United States
    Contribution
    Formal analysis, Methodology
    Competing interests
    No competing interests declared
  15. Jennifer H Gutzman

    Department of Biological Sciences, Cell and Molecular Biology, University of Wisconsin Milwaukee, Milwaukee, United States
    Contribution
    Resources, Funding acquisition, Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7725-6923
  16. Dylan T Burnette

    Department of Cell and Developmental Biology, Vanderbilt University, Nashville, United States
    Contribution
    Conceptualization, Resources, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    For correspondence
    dylan.burnette@vanderbilt.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2571-7038

Funding

National Heart, Lung, and Blood Institute (F31 HL136081)

  • Aidan M Fenix

American Heart Association (16PRE29100014)

  • Aidan M Fenix

National Heart, Lung, and Blood Institute (K08 HL116803)

  • Jason R Becker

National Heart, Lung, and Blood Institute (RO1 HL037675)

  • David M Bader

University of Wisconsin-Milwaukee (ResearchGrowth Initiative)

  • Jennifer H Gutzman

National Institute of General Medical Sciences (R35 GM125028)

  • Dylan Tyler Burnette

National Cancer Institute (P50 CA095103)

  • Dylan Tyler Burnette

American Heart Association (17SDG33460353)

  • Dylan Tyler Burnette

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

Acknowledgements

We thank the M Tyska lab and the Epithelial Biology Center (EBC) at Vanderbilt University for invaluable discussions and reagents. Janice Williams of the Cell Imaging Shared Resources was instrumental in acquiring the EM images in Figure 1A. We give a special thanks to Sean Schaffer and Bryan Millis of the Cell Imaging Shared Resources and Nikon Center of Excellence at Vanderbilt University for help with and maintenanc of the SIM and spinning disk microscopes, and Anthony Tharp and Josh Luffman for the CDB CORE Equipment. The authors declare no competing financial interests.

This work was supported by Vanderbilt University School of Medicine Molecular Biophysics Training Grant T32 GM08320 to AMF, an American Heart Association pre-doctoral fellowship #16PRE29100014 to AMF, a NIH F31 pre-doctoral fellowship F31 HL136081 to AMF, a Career Development Award from the National Cancer Institute SPORE in GI Cancer P50 CA095103 to DTB, an American Heart Association Scientist Development Grant #17SDG33460353 to DTB a MIRA Grant R35 GM125028 from NIGMS to DTB, RO1 HL037675 from NHLBI to DMB, a K08 HL116803 to JRB, and an University of Wisconsin Research Growth Initiative grant to JHG.

Senior and Reviewing Editor

  1. Anna Akhmanova, Utrecht University, Netherlands

Publication history

  1. Received: September 18, 2018
  2. Accepted: December 11, 2018
  3. Accepted Manuscript published: December 12, 2018 (version 1)
  4. Version of Record published: December 27, 2018 (version 2)

Copyright

© 2018, Fenix 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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