RESEARCH ARTICLE 3827

Development 139, 3827-3837 (2012) doi:10.1242/dev.079178© 2012. Published by The Company of Biologists Ltd

INTRODUCTIONThe size, shape and organization of tissues and individual cellsdirectly impact their physiology. A pressing question in cell anddevelopmental biology is how different morphogenetic processesthat are necessary for the functional development of a commontissue are coordinated. Skeletal muscle provides an ideal systemwith which to study structure-function relationships as it is highlyorganized with a physiologically testable output.

The cellular unit of all metazoan muscle is the myofiber, asyncytial cell that is formed from the iterative fusion of myoblasts.The correlation between structure and function is highlighted bythe organization of myosin and actin within the sarcomere thatprovides the contractile apparatus (Squire, 1997). However, otheraspects of skeletal muscle structure, including myofiber size andorganelle positioning, are also crucial for its function.

Myofiber size encompasses a variety of measurements,including length, width, depth and volume, and size is importantfor muscle output, as smaller myofibers provide less contractileforce than larger muscles (Fitts et al., 1991). During muscleformation in Drosophila, muscle cell growth occurs in threedistinct phases (Volk, 1999; Schnorrer and Dickson, 2004). Asfusion begins, the muscle cell breaks symmetry, adds mass, andelongates to form polarized structures with a long axis and a shortaxis. Once the long axis is established, the muscle cell extendsfilopodia at each end in search of an attachment with a tendon cell.Finally, the filopodial protrusions stop, the muscle cell polesbecome smooth, and stable muscle-tendon attachments are made.In Drosophila embryos, muscle size has often been correlated with

the number of fusion events (Bate, 1990), but it is unlikely thatfusion alone determines myofiber size. In Drosophila larvae and inmammals, a number of signaling pathways have been demonstratedto control muscle size through growth regulation (Demontis andPerrimon, 2009; Rommel et al., 2001). Although several guidancecues and signaling mechanisms are known to regulate the directionof muscle growth (Schejter and Baylies, 2010; Schweitzer et al.,2010), the factors and mechanisms that directly provide thenecessary forces that promote oriented and regulated musclegrowth during early myogenesis are not known.

The importance of organelle positioning is highlighted by thelocalization of the myonuclei. In skeletal muscle, nuclei resideabove the sarcomere at the periphery of the muscle fiber and arepositioned to maximize their internuclear distance. Moreover,mispositioned myonuclei, in conjunction with smaller myofibers,are a hallmark of many muscle disorders and have been useddiagnostically for decades (Pierson et al., 2007; Romero, 2010).The mechanisms by which nuclei are moved throughout the muscle(Metzger et al., 2012) and anchored at the neuromuscular junction(NMJ) (Grady et al., 2005; Lei et al., 2009) are beginning toemerge; however, only a small number of factors necessary forthese processes have been identified. Additionally, potential linksbetween muscle size and myonuclear position have not beenexplored. In other systems, the microtubule cytoskeleton regulatesboth polarized growth and nuclear position. Cytoplasmic Dynein(Dynein) is a crucial factor in both processes. During nuclearmovement in the first cell division of C. elegans and in migratingneurons, cortically anchored Dynein pulls microtubule minus endsand the attached nuclei towards itself (Gönczy, 2002; Tsai et al.,2007). During polarized cell growth, Dynein contributes to theestablishment of the polarity axis (Etienne-Manneville and Hall,2001; Palazzo et al., 2001) that is necessary to localize the factorsfor cell protrusion (Prigozhina and Waterman-Storer, 2004;Schmoranzer et al., 2003). Given these functions, Dynein maycontribute to both myofiber size and myonuclear positioning andmight integrate the two processes.

1Program in Developmental Biology, Sloan Kettering Institute, New York, NY 10065,USA. 2Cell and Developmental Biology, Weill Cornell Graduate School of MedicalSciences, Cornell University, New York, NY 10065, USA.

*Author for correspondence (m-baylies@ski.mskcc.org)

Accepted 16 July 2012

SUMMARYVarious muscle diseases present with aberrant muscle cell morphologies characterized by smaller myofibers with mispositioned nuclei.The mechanisms that normally control these processes, whether they are linked, and their contribution to muscle weakness indisease, are not known. We examined the role of Dynein and Dynein-interacting proteins during Drosophila muscle developmentand found that several factors, including Dynein heavy chain, Dynein light chain and Partner of inscuteable, contribute to theregulation of both muscle length and myonuclear positioning. However, Lis1 contributes only to Dynein-dependent muscle lengthdetermination, whereas CLIP-190 and Glued contribute only to Dynein-dependent myonuclear positioning. Mechanistically,microtubule density at muscle poles is decreased in CLIP-190 mutants, suggesting that microtubule-cortex interactions facilitatemyonuclear positioning. In Lis1 mutants, Dynein hyperaccumulates at the muscle poles with a sharper localization pattern, suggestingthat retrograde trafficking contributes to muscle length. Both Lis1 and CLIP-190 act downstream of Dynein accumulation at thecortex, suggesting that they specify Dynein function within a single location. Finally, defects in muscle length or myonuclearpositioning correlate with impaired muscle function in vivo, suggesting that both processes are essential for muscle function.

KEY WORDS: Drosophila, Dynein, Nuclear positioning, Muscle size

Muscle length and myonuclear position are independentlyregulated by distinct Dynein pathwaysEric S. Folker1, Victoria K. Schulman1,2 and Mary K. Baylies1,2,*

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To investigate how myofiber size and myonuclear positioningare regulated, and whether the two processes are co-dependent, wehave studied the role of Dynein during muscle morphogenesis inDrosophila. We establish muscle length, which is an aspect ofmuscle size, and myonuclear positioning as independent aspects ofmuscle morphogenesis. We further show that the microtubulecytoskeleton contributes to both processes. Specifically, Dyneinregulates muscle length and myonuclear positioning withoverlapping, but distinct, sets of proteins. The Dynein heavy chain(Dhc64C), Dynein light chain (Dlc90F; Tctex1 in mammals) andPartner of inscuteable [Pins; Rapsynoid (Raps) – FlyBase] arenecessary regulators of both muscle length and myonuclearpositioning. Subsequently, the two pathways become distinct:muscle length is regulated by a Dynein-Lis1-dependentmechanism, whereas myonuclear positioning is regulated by aDynein–CLIP-190–Glued-dependent mechanism.

Mechanistically, in mutant embryos that lack Dynein (Dhc64C4-19) or fail to localize Dynein to the muscle pole (raps193),both muscle length and myonuclear positioning are affected,suggesting that Dynein activity at the muscle pole is required forboth processes. However, Dynein localizes to the muscle pole inboth Lis1 and CLIP-190 mutant embryos, indicating that Dynein-dependent regulation of muscle length and Dynein-dependentregulation of myonuclear positioning are specified downstream ofits arrival at the muscle pole. Lis1 mutant embryos exhibithyperaccumulation of Dynein at the muscle pole with a tighterfocus of peak intensity, suggesting that Lis1-dependent activationof Dynein retrograde trafficking is essential for muscles to achievetheir proper length. CLIP-190 mutant embryos have fewermicrotubules near the myofiber poles despite normal Dyneinlocalization, suggesting that CLIP-190 promotes microtubule-cellcortex interactions that are necessary for Dynein-dependentmyonuclear positioning. Together, these data suggest that bothCLIP-190 and Lis1 work downstream of Dynein localization to themuscle pole and thus specify Dynein function within this singlelocation. Differential regulation of Dynein activity within a singlelocation is novel and likely to be important during morphogeneticprocesses for which proper timing is essential. Finally, both musclelength and myonuclear positioning are important for musclefunction, as defects in either process correlate with larvae thatcrawl more slowly than controls.

MATERIALS AND METHODSDrosophila geneticsStocks were grown under standard conditions. Stocks used were apME-NLS::dsRed (Richardson et al., 2007), Dhc64C4-19 and Dhc64C6-10 (Gepneret al., 1996), UAS-Dhc64C-RNAi (Vienna Drosophila RNAi Center,v28053), dlc90F05089 (Caggese et al., 2001), lis1K11702 (Lei and Warrior,2000), lis1G10.14 (Liu et al., 1999), UAS-Lis1-RNAi (Vienna DrosophilaRNAi Center, v6216), clip190KG06490 (Bloomington Drosophila StockCenter, 14493), UAS-Clip190-RNAi (Vienna Drosophila RNAi Center,v107176), raps193 (Parmentier et al., 2000), inscP49 (Kraut and Campos-Ortega, 1996) and UAS-Glued-RNAi (Vienna Drosophila RNAi Center,v3785). Mutants were balanced and identified using CTG (CyO, twi-Gal4,UAS-2xeGFP), TTG (TM3, twi-Gal4, UAS-2xeGFP), TM6B, CyOP[w+wgen11lacZ] and TM3 Sb1 Dfd-lacZ.

ImmunohistochemistryEmbryos were collected at 25°C and fixed with 4% EM-gradeparaformaldehyde (Polysciences, 00380) diluted in PBS (Roche,11666789001) and an equal volume of heptane to allow permeabilization.In all cases, embryos were devitellinized by vortexing in a 1:1methanol:heptane solution. Larvae were dissected in ice-cold HL3.1 aspreviously described (Brent et al., 2009) and fixed with 10% formalin

(Sigma, HT501128-4L). Embryos and larvae were mounted in ProLongGold (Invitrogen) for fluorescent stainings. Antibodies were preabsorbedwhere noted (PA) and used at the following final dilutions: rabbit anti-dsRed (PA, 1:400, Clontech 632496), rat anti-Tropomyosin (PA, 1:500,Abcam ab50567), mouse anti-GFP (PA, 1:200, Clontech 632381), mouseanti-DHC (1:50, Developmental Studies Hybridoma Bank), mouse anti-Tubulin (1:500, Sigma T9026), chicken anti--galactosidase (PA, 1:1000,Novus Biologicals NB100-2045), mouse anti--PS-Integrin (1:100,Developmental Studies Hybridoma Bank) and mouse anti-Discs large(1:200, Developmental Studies Hybridoma Bank). We used Alexa Fluor488-, Alexa Fluor 555- and Alexa Fluor 647-conjugated fluorescentsecondary antibodies (1:200), Alexa Fluor 647-conjugated phalloidin(1:100) and Hoechst 33342 (1 g/ml) for fluorescent stains (all Invitrogen).Fluorescent images were acquired on a Leica SP5 laser-scanning confocalmicroscope equipped with a 63� 1.4 NA HCX PL Apochromat oilobjective and LAS AF 2.2 software. Images were processed using AdobePhotoshop CS4. Maximum intensity projections of confocal z-stacks wererendered using Volocity Visualization software (Improvision).

Nuclear position and muscle length measurementsEmbryos were staged based on standard laboratory procedures: overallembryo shape, the intensity of the apRed and Tropomyosin signals, gutmorphology, and the morphology of the trachea (Beckett and Baylies,2007). Measurements were taken from confocal projections of embryosand acquired using the Line function of ImageJ software (NIH). For eachgenotype, all four lateral transverse (LT) muscles were measured in threehemisegments from each of 20 embryos from four independentexperiments. Three measurements were taken within each LT muscle: (1)myotube length and (2, 3) the shortest distance between the myotube polesand the outermost edge of the nearest nucleus (supplementary material Fig.S1). The edges of the nuclei and ends of the myotubes were defined by theclear change from signal to background within the relevant viewingchannel. Similar measurements were performed using -PS-Integrin toidentify the ends of the LT muscles. The statistical significance ofdifferences in measurements was assessed using a Student’s t-test tocompare each experimental genotype with wild-type apRed controls.Statistical analysis was performed with Prism 4.0 (GraphPad).

Because the live stage 17 embryos offer few markers for accuratestaging, staging was achieved by timed lays: after a pre-lay period,embryos were collected for 15 minutes and then aged for 16.5 hours.Embryos were then dechorionated and mounted on coverslips inhalocarbon oil and images were acquired on a Zeiss Axioplan 2 microscopewith a 20� PlanNeoFluor 0.50 NA objective and an Axiocam MRmcamera. Nuclear spread measurements were performed using the Linefunction in ImageJ to measure the distance between the dorsal edge of thedorsal nucleus and the ventral edge of the ventral nucleus. Additionally,these same images of stage 17 embryos were used to count the number ofnuclei per hemisegment, which was used as a measure of the number ofmyoblast fusion events.

Microtubule organization measurementsConfocal z-stacks were sequentially acquired to maximize the signal andresolution at the myofiber poles and to minimize interference from theinternal muscles and the epidermis. Images were acquired with a 63� 1.4NA HCX PL Apochromat oil objective and a 10� optical zoom. z-stacksthrough a depth of 2 m with a step size of 0.25 m were acquired for allanalyzed images. z-stacks were then assembled into confocal projectionsusing Volocity, which were then exported to ImageJ for analysis. Analysiswas performed on 20 embryos from four independent experiments. Tocount the number of microtubules in contact with the muscle pole, a 2�2m box was drawn from the tip of the muscle toward the center and thenumber of microtubules in this region were counted.

To determine the average intensity, the Measure Average IntensityFunction in ImageJ was performed on the distal 3 m of the myofiber. Theintensity of Tubulin staining was standardized against the intensity ofTropomyosin staining to control for sample variation. For each genotype,all four LT muscles were measured in three hemisegments from each of 20embryos from four independent experiments. Student’s t-test was used to

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compare each experimental genotype with wild-type apRed controls.Statistical analysis was performed with Prism 4.0.

Dynein localization measurementsEmbryos were collected and fixed with 10% formalin diluted 1:1 inheptane for 20 minutes, then rinsed three times in PBS containing 0.6%Triton X-100, then fixed with 4% paraformaldehyde diluted 1:1 in heptanefor 20 minutes. Antibody incubations were performed in PBSsupplemented with 1% BSA and 0.6% Triton X-100. Image acquisition andprocessing were as for microtubule quantification. Dyneinimmunofluorescence intensity was assessed by linescan analysis usingImageJ. Positions of linescans were chosen randomly on the Tropomyosinimage and the line then copied to the identical position on the Dyneinimage. The intensity of Dynein immunofluorescence at each point wasmeasured relative to Tropomyosin intensity at that same point to controlfor sample variation, and the ratios were multiplied by 100. Threehemisegments from 20 embryos from four independent experiments wereanalyzed. The greatest pixel intensity from each embryo was used todetermine the peak intensity. To determine the width of peak intensity, thepixels that were ≥80% of peak intensity were determined and the distancebetween the first pixel and the last pixel that fitted these criteria wasmeasured. For each genotype, all four LT muscles were measured in threehemisegments from each of five embryos. Student’s t-test was used tocompare each experimental genotype with wild-type apRed controls.Statistical analysis was performed with Prism 4.0. The heat maprepresentation of Dynein immunofluorescence was generated using theCool application of the standard Lookup tools in ImageJ.

Larval behaviorLarval speed was assessed as previously described (Louis et al., 2008;Metzger et al., 2012) with minor modifications. Stage 16 and 17 embryoswere selected for the absence of the fluorescent balancer and placed onyeast-coated apple juice agar plates at 22°C overnight. L1 larvae wereselected the following day and placed into vials of standard fly foodcontaining Bromophenol Blue (Bio-Rad, 161-0404). L3 larvae were pickedfrom the vial 6 days later and individually tracked as they crawled towardsan odor source of ethyl butyrate (3.3%; Sigma, E15701) diluted in paraffinoil (Fluka, 76235). Their movements were recorded with a CCD camerafor either five total minutes or until they reached the odor source or thewalls of the apparatus. The tracks were processed using Ethovisionsoftware (Nodlus). At least 20 larvae were tracked for each genotype.Tracked larvae were dissected, stained and analyzed as follows.Internuclear distance was measured in ImageJ by drawing lines betweenthe center of each nucleus and its nearest neighbor using projection images.Muscle size was measured by determining the area of confocal projectionsof each muscle. To calculate the internuclear distance as a function ofmuscle size, the average internuclear distance for a given muscle wasdivided by the area of that specific muscle. Nine muscles from three larvaewere used for quantification.

RESULTSDynein regulates muscle length and myonuclearpositionEach body wall muscle in the Drosophila embryo and larvaconsists of a single syncytial myofiber similar to the individualmyofibers in mammalian muscle. To assess the role of Dyneinduring Drosophila muscle morphogenesis, the distribution of nucleiwithin the lateral transverse (LT) muscles was examined in wild-type embryos and in embryos in which Dynein was zygoticallyremoved with either the Dhc64C4-19 (null) or the Dhc64C6-10

(hypomorphic) allele (Gepner et al., 1996). The myonuclei in theLT muscles were visualized by expression of the apterousmesodermal enhancer fused to a nuclear localization signal and thefluorescent protein DsRed (apRed) (Richardson et al., 2007), whichspecifically labels the nuclei of the LT muscles. Using this reporter,live stage 17 embryos [16.5-20 hours after egg lay (AEL)] were

examined. Dhc64C mutant embryos had the same number ofapRed nuclei per hemisegment as controls, indicating that myoblastfusion was unaffected (Fig. 1A,B). Although the nuclei werealigned in columns within each LT muscle in embryos of eachgenotype, the distance between the most ventral and the mostdorsal nuclei was reduced in Dhc64C mutant embryos (Fig. 1A,C).To determine whether the difference in the distribution of nucleiwas caused by defective nuclear positioning or effects on musclesize, we examined late stage 16 embryos (16 hours AEL), whichare amenable to whole-mount immunostaining.

The general muscle pattern as revealed by Tropomyosin staining(Fig. 1D) was normal in Dhc64C mutant embryos. Additionally, -PS-Integrin staining was normal. Thus, all muscles were present,properly oriented and attached to tendon cells, indicating thatspecification, guidance and attachment mechanisms are functional.In late stage 16 (16 hours AEL) control embryos, the myonucleiwere in two clusters within each LT muscle: one cluster at thedorsal pole and the other at the ventral pole (Fig. 1D;supplementary material Fig. S1) (Metzger et al., 2012). In embryoshomozygous for either Dhc64C allele, the nuclei were also in twoclusters; however, the clusters were mispositioned 50% furtherfrom the muscle poles than in wild-type and heterozygous controls(Fig. 1D,E; supplementary material Fig. S2). Additionally, thelength of the LT muscles was reduced by 15% in embryoshomozygous for either Dhc64C allele as compared with wild-typeand heterozygous controls (Fig. 1D,F; supplementary material Fig.S2). To account for the difference in muscle length, the distancefrom each cluster of nuclei to the nearest muscle pole wascalculated as a percentage of muscle length (Fig. 1G). Thismeasurement was used with respect to all subsequent genotypes.To validate the use of Tropomyosin as a marker for muscle ends,we performed identical measurements using -PS-Integrin to markthe muscle ends. These analyses produced identical results (Fig.1D; supplementary material Fig. S2C,D).

To confirm that the effects of Dynein were autonomous tomuscle, we used the GAL4/UAS system to deplete Dyneinspecifically in the mesoderm using twist-GAL4 to express UAS-Dhc64C-RNAi (Dietzl et al., 2007). Tissue-specific depletion ofDhc64C recapitulated the effects of the Dhc64C alleles: the LTmuscles were 12% shorter than controls and the nuclei werepositioned 40% further from the muscle poles than in controls (Fig.1D-G). These data indicate that Dynein is specifically required inthe muscle to generate muscles of proper length and to properlyposition myonuclei.

Given that myonuclear position seems to be affected similarly instage 16 and stage 17 embryos, we examined embryos at earlierstages in their development to determine when the defects inmuscle length and myonuclear positioning become evident. UsingTropomyosin to identify the muscles and a combination of tracheamorphology and gut morphology to assess embryo age, wedetermined the length of muscles and the positioning of myonucleiin late stage 14 (11 hours AEL), stage 15 (13 hours AEL) and stage16 (16 hours AEL) embryos (Fig. 2). At stage 14, the length of themuscles in control and Dhc64C4-19 embryos was similar. However,by stage 15 and through stage 16 the muscles in the Dhc64C4-19

embryos were significantly shorter (17% and 14%, respectively)than those in the control embryos.

The effects on myonuclear positioning at the different stageswere more complex. The nuclei were 30% further from the musclepoles in late stage 14 Dhc64C4-19 embryos than in controls (Fig.2C); however, only the positioning relative to the dorsal musclepole was statistically significant. At stage 15, myonuclei were

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similarly positioned in control and Dhc64C4-19 embryos (Fig. 2D).In stage 16 embryos, the myonuclei were significantlymispositioned (50% further from the poles) at both the dorsal andventral poles of the muscle in Dhc64C4-19 mutant embryos relativeto controls (Fig. 2E).

These data indicate that the defects in muscle length arise priorto muscle-tendon attachment and, together with the muscle-specificdepletion of Dhc64C, suggest that the effects are muscleautonomous and not due to defects in tendon specification.

Dynein-interacting proteins contribute to Dynein-dependent regulation of muscle length andmyonuclear positioningTo determine whether the roles of Dynein in specifying musclelength and myonuclear positioning are functionally coupled, wetested known Dynein-interacting proteins for their contributions toeach process. We focused on factors that are general regulators ofDynein activity or have been found to specifically contribute toDynein-mediated nuclear movements in other systems. Weexamined Dynein light chains, which can regulate Dynein activityboth positively (Sönnichsen et al., 2005) and negatively (O’Rourkeet al., 2007); Glued, which increases Dynein motor processivityand mediates Dynein-cargo interactions (Gill et al., 1991; Vaughanet al., 2002; Waterman-Storer et al., 1997); Pins, which anchorsDynein at the cortex to move nuclei in other systems (Gotta et al.,2003; Hughes et al., 2004); Lis1, which maintains Dynein in anactive state and has been implicated in nuclear movements in yeast

and neurons (McKenney et al., 2010; Mesngon et al., 2006;Sheeman et al., 2003); and CLIP-190, which works with Lis1 toregulate Dynein activity (Tanenbaum et al., 2008).

We tested two Dynein light chains, Cdlc2 and Dlc90F, which arethe Drosophila orthologs of Dynein light chain 1/2 and Tctex1,respectively (Caggese et al., 2001; Goldstein and Gunawardena,2000), by RNAi-mediated depletion (Dietzl et al., 2007). Depletionof Cdlc2 specifically in mesoderm and muscle did not affectmuscle length or myonuclear positioning, whereas depletion ofDlc90F resulted in shorter muscles with mispositioned nuclei.Additionally, embryos that were homozygous for dlc90F05089

(Caggese et al., 2001) or the pins allele raps193 (Parmentier et al.,2000), produced muscles that were 12% shorter with nuclei thatwere 40-50% further from the muscle poles than in wild-type andheterozygous control embryos (Fig. 3A-C; supplementary materialFig. S2). These data indicate that Dlc90F and Pins contribute toboth muscle length and myonuclear positioning.

Embryos homozygous for either of two Lis1 alleles, lis1K11702

(Lei and Warrior, 2000) and lis1G10.14 (Liu et al., 1999), producedmuscles that were 15% shorter than those of wild-type andheterozygous controls. However, myonuclear positioning wasnormal in Lis1 homozygous embryos. Conversely, embryos thatwere depleted of Glued (Dietzl et al., 2007) in muscle and embryosthat were homozygous for clip190KG06490 (Bellen et al., 2004)produced muscles of similar length to controls. However, in theseembryos the nuclei were positioned 50% further from the musclepoles than in wild-type and heterozygous controls (Fig. 3A-C;

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Fig. 1. Cytoplasmic Dynein regulates muscle length andmyonuclear position. (A)Fluorescence images of apRed nucleiin the lateral transverse (LT) muscles of live stage 17 Drosophilaembryos (16.5-20 hours AEL) of the indicated genotypes. (B)Thenumber of nuclei per hemisegment in stage 17 embryos of theindicated genotypes. (C)The distance between the dorsalmostand ventralmost nuclei in stage 17 embryos of the indicatedgenotypes. (D)Immunofluorescence images of stage 16 (16hours AEL) embryos of the genotypes noted to the left andantigen listed at the top of the first image. Green, Tropomyosin(muscle); red, DsRed (nuclei); blue, -PS-Integrin in merge.Arrows in the control panels denote points from whichmeasurements were made for all genotypes. The distancebetween the points indicated by the yellow arrows in theTropomyosin and -PS-Integrin panels was used to determinemuscle length. The distance between the pair of gray arrows atthe top of the merged image was used to determine thedistance between the nuclei and the dorsal pole. The distancebetween the pair of white arrows at the bottom of the mergedimage was used to determine the distance between the nucleiand the ventral pole. (E)The shortest distance between theindicated pole of the LT muscles (gray, dorsal; white, ventral) andthe nearest cluster of nuclei in stage 16 embryos. (F)LT musclelength in stage 16 embryos. (G)The shortest distance betweenthe indicated pole of the LT muscles (gray, dorsal; white, ventral)and the nearest cluster of nuclei normalized for muscle length instage 16 embryos of the indicated genotypes. Error bars indicates.d.; **P<0.01, *P<0.05. Scale bars: 10m.

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supplementary material Fig. S2). The effects seen on myonuclearpositioning in the clip190KG06490 homozygote were phenocopied inembryos that were transheterozygous for clip190KG06490 and eachof three different deficiencies that removed CLIP-190:

Df(2L)BSC294, Df(2L)Exel7068 and Df(2L)Exel8036(supplementary material Fig. S2). Additionally, we measured theventral longitudinal muscle VL3 and found that its length in theembryo was similar in each genetic background (supplementarymaterial Fig. S2). These data suggest that a unique feature of theLT muscles makes them more susceptible to Dynein levels and/oractivity with respect to muscle growth in the embryo.

The number and position of nuclei in stage 17 embryos were alsoexamined. Embryos from each genotype had the same number ofapRed nuclei per hemisegment, indicating that the defects in musclelength did not result from impaired fusion (Fig. 1B). The clusters ofnuclei in each genotype did resolve into columns during stage 17, butthe distance between the ventral and dorsal nuclei was reduced by15% compared with controls. Moreover, the distance between themost distal nuclei was similar in stage 16 and stage 17 embryos foreach genotype (supplementary material Fig. S3).

To determine whether the Dynein-interacting proteins work with,or independently of, Dynein, we examined embryos that were doublyheterozygous for Dhc64C mutations and mutations in each of theassociated genes. In these, and all subsequent genetic interactions,maternal effects were controlled for by providing each allele fromthe mother or the father in reciprocal crosses. Both Dhc64C4-19 andDhc64C6-10 were used in combination with alleles for each putativeDynein-interacting gene. Double heterozygotes of dlc90F05089 andDhc64C4-19 or of dlc90F05089 and Dhc64C6-10 produced muscles thatwere 15% shorter than controls and with their nuclei 50% furtherfrom the muscle poles (Fig. 4A-C). These data indicate that Dlc90F(Tctex1) regulates Dynein activity for both muscle length andmyonuclear positioning. All Lis1–/+; Dhc64C–/+ doublyheterozygous embryos produced muscles that were 15% shorter thanthose in control embryos, but with properly positioned myonuclei atthe muscle poles (Fig. 4A-C). Conversely, CLIP-190–/+; Dhc64C–/+doubly heterozygous embryos produced muscles that were the samelength as those of control embryos but with nuclei 40% further fromthe muscle poles (Fig. 4A-C; supplementary material Fig. S3).Dhc64C–, +/+, raps193 doubly heterozygous embryos and embryos

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Fig. 3. Dynein-interacting proteinsregulate myotube length and/ormyonuclear position.(A)Immunofluorescence images ofstage 16 Drosophila embryos of theindicated genotypes. Green,Tropomyosin; red, DsRed; blue, -PS-Integrin in merge. The antigen forgrayscale images is listed at the top ofthe first image. Scale bar: 10m. (B)Thelength of the LT muscles in stage 16embryos. (C)The shortest distancebetween the indicated pole of the LTmuscles (gray, dorsal; white, ventral) andthe nearest cluster of nuclei normalizedfor muscle length. Error bars indicates.d.; *P<0.05, **P<0.01.

Fig. 2. Stage-dependent effects of Dynein on muscle length andmyonuclear positioning. (A)Immunofluorescence images of themuscles (green, Tropomyosin) and nuclei (red, DsRed) at stage 14, 15and 16 in control and Dhc64C4-19 Drosophila embryos. Scale bar:10m. (B)The length of the muscles in control (gray) and Dhc64C4-19

(black) embryos at stage 14, 15 and 16. (C-E)The distance between thenearest nucleus and either the dorsal (gray) or ventral (white) pole ofthe muscle in stage 14 (C), 15 (D) and 16 (E) embryos. Error barsindicate s.d.; *P<0.05, **P<0.01.

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depleted of Glued in the Dhc64C–/+ background died prior to stage16 and could not be evaluated. These data indicate that Lis1 andCLIP-190 regulate muscle length and myonuclear position,respectively, via interactions with Dynein, and not by Dynein-independent mechanisms.

The two Dynein-dependent pathways areseparableTo confirm that Dynein-dependent myonuclear positioning andDynein-dependent determination of muscle length are regulated byseparate mechanisms, we examined the interactions betweenDynein regulatory proteins. Both Lis1 and CLIP-190 functionally

interacted with dlc90F05089, and these interactions recapitulatedtheir interactions with Dhc64C: lis1K11702/+; dlc90F05089/+ andlis1G10.14/+; dlc90F05089/+ double heterozygotes had muscles thatwere 20% shorter than those of control embryos, whereasclip190KG06490/+; dlc90F05089/+ double heterozygotes had musclesin which the nuclei were mispositioned, 30% further from themuscle poles compared with controls. Similarly, pins functionallyinteracted with both Lis1 and CLIP-190: lis1K11702/+; raps193/+double heterozygotes had muscles that were 20% shorter than thoseof control embryos and clip190KG06490/+; raps193/+ doubleheterozygotes had normal sized muscles with mispositionedmyonuclei that were 60% further from the muscle poles compared

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Fig. 4. Interactions between Dynein and associated genes during muscle development. (A)Immunofluorescence images of stage 16Drosophila embryos that are doubly heterozygous for Dhc64C4-19 and the indicated allele. Green, Tropomyosin; red, DsRed. Scale bar: 10m. (B)LTmuscle length in stage 16 embryos that are doubly heterozygous for the Dhc64C allele shown beneath the histogram and the listed alleles. (C)Theshortest distance between the indicated pole of the LT muscles (gray, dorsal; white, ventral) and the nearest cluster of nuclei normalized for musclelength in stage 16 embryos that are doubly heterozygous for the Dhc64C allele indicated beneath the histogram and the listed alleles. Error barsindicate s.d.; **P<0.01.

Fig. 5. The pathways controlling muscle length and myonuclear position do not genetically interact. (A)Immunofluorescence images ofstage 16 Drosophila embryos that are doubly heterozygous for the indicated alleles. Green, Tropomyosin; red, DsRed. Scale bar: 10m. (B)LTmuscle length in stage 16 embryos that are doubly heterozygous for the indicated alleles. (C)The shortest distance between the indicated pole ofthe LT muscles (gray, dorsal; white, ventral) and the nearest cluster of nuclei in stage 16 embryos normalized for muscle length. Error bars indicates.d.; *P<0.05, **P<0.01. D

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with controls (Fig. 5A-C; supplementary material Fig. S3). Theinteraction of dlc90F05089 with pins was also examined but resultedin embryonic death prior to stage 16 and could not be evaluated.

Lis1 contributes only to muscle length, whereas CLIP-190contributes solely to myonuclear positioning. We tested whetherLis1 functionally interacts with CLIP-190. Lis1–, +/+,clip190KG06490 double heterozygotes were similar to controlembryos in both muscle length and myonuclear positioning. Thisindicates that Dynein-dependent muscle length and Dynein-dependent myonuclear positioning are indeed mechanisticallydistinct processes (Fig. 5A-C; supplementary material Fig. S3).

Lis1 affects Dynein localization and CLIP-190affects microtubule-cortex interactionsWe next examined the effects of Dhc64C, Lis1, CLIP-190 and rapsmutants on Dynein localization (Fig. 6A) and microtubuleorganization (Fig. 7A) to understand how Dynein differentiallyregulates muscle length and myonuclear positioning. Usingconfocal projections, linescans of Dynein immunofluorescenceintensity (Fig. 6B) were employed to measure Dynein localization.To control for sample variation, the intensity of Dyneinimmunofluorescence was measured relative to the intensity ofTropomyosin immunofluorescence at the same point. Both the peakof Dynein immunofluorescence (Fig. 6C) and width of peakimmunofluorescence intensity (Fig. 6D) were quantified.clip190KG06490 did not significantly affect Dynein localization. Lis1and raps mutants did affect Dynein localization, althoughdifferently. In raps193 mutant embryos, the peak intensity of Dyneinimmunofluorescence was decreased compared with controls,whereas the width of peak intensity was increased (Fig. 6A-D),suggesting that Pins is required for Dynein localization to the

muscle poles. In Lis1 mutant embryos, the peak intensity of Dyneinimmunofluorescence was increased compared with controls and thewidth of peak intensity was slightly decreased (Fig. 6A-D). Thehigher peak intensity combined with the sharper localizationsuggested that Lis1 does not contribute to the initial localization ofDynein to the muscle pole, but rather to the retrograde traffickingof Dynein away from the muscle pole. Together, these data indicatethat proper muscle length requires Dynein localization to themuscle pole, and that raps193 results in shorter muscles due to pooraccumulation of Dynein at the myofiber pole. Additionally, tightlyfocused hyperaccumulation of Dynein, as seen in Lis1 mutantembryos, also affects muscle length, suggesting that trafficking ofDynein away from the myofiber pole is mediated by Lis1 and isnecessary to generate muscles of the proper length.

Similarly, confocal projections of embryos immunostained forTubulin and Tropomyosin were used to assess the presence ofmicrotubules near myofiber poles (Fig. 7A). Two differentmeasures – the number of microtubules in contact with themyofiber pole (Fig. 7B) and the average intensity of tubulinimmunofluorescence in the distal 3 m of the myofiber (Fig. 7C)– indicated that only clip190KG06490 homozygotes had significantlyfewer microtubules near the muscle pole than all other genotypes.These data suggest that CLIP-190 is necessary to mediateinteractions between microtubules and the myofiber cortex that arethen utilized by Dynein to move nuclei towards the muscle pole.

Defects in muscle size and myonuclear positionimpair muscle functionLarval crawling towards an odorant stimulus was used to assess theimportance of Dynein-dependent muscle size regulation andDynein-dependent myonuclear positioning to muscle physiology.

3833RESEARCH ARTICLEDynein-dependent muscle formation

Fig. 6. Dynein hyperaccumulates inLis1 mutant embryos. (A)Confocalprojections of a single hemisegmentfrom stage 16 Drosophila embryosimmunostained for Tropomyosin(green) and Dynein heavy chain (red).The boxed regions are shown at highermagnification to the right, withTropomyosin shown in grayscale andDynein shown as a heat map (the scaleindicates relative intensity). The regionsoutlined by the green dotted lineswere used for linescan analysis. Scalebars: left-hand panel, 5m; right-handpanel, 10m. (B)Representativelinescan analysis indicating theintensity of Dynein heavy chainimmunofluorescence as a function ofposition across the muscle pole.(C)The peak intensity signal for Dyneinheavy chain immunofluorescence inthe indicated genotypes. (D)The widthof the peak intensity of Dynein heavychain immunofluorescence. Error barsindicate s.d.; *P<0.05, **P<0.01,compared with control.

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Each genotype tested crawled towards the stimulus, indicating thatall genotypes perceived and responded to the stimulus. However,L3 larvae that were homozygous for clip190KG06940 or lis1G10.14

crawled towards the stimulus 33% more slowly than controls (Fig.8A). Dhc64C4-19, Dhc64C6-10 and lis1K11702 were early larval lethal,thus their locomotive ability could not be tested. However, whenDhc64C was depleted during embryonic and larval developmentby driving the UAS-Dhc64C-RNAi construct with the muscle-specific driver DMef2-Gal4, these larvae also crawled towards thestimulus 40% more slowly than controls (Fig. 8A,B). Additionally,muscle-specific depletion of CLIP-190 and Lis1 inhibited larvallocomotion by 20% (Fig. 8A,B). That the depletion was musclespecific confirms that the effects on crawling are muscleautonomous. Furthermore, the NMJ, as measured by boutonnumber and distribution, was similar to that of the control in eachof the genotypes tested (supplementary material Fig. S4). Finally,lis1G10.14, +/+, clip190KG06490 doubly heterozygous larvae crawledtowards the stimulus at the same speed as control larvae (Fig.8A,B), further indicating the independence of these two geneticpathways.

The larvae were then dissected and their musculature wasexamined. The internuclear distance was reduced by 50-60% inlis1G10.14 and clip190KG06490 larvae and in muscle depleted ofDhc64C, Lis1 or CLIP-190 (Fig. 8C,D). However, the muscles inlis1G10.14, Lis1-depleted and Dhc64C-depleted larvae were 35%smaller than in control and clip190KG06490 larvae (Fig. 8E). Becausethe muscle size was variable between genotypes, internucleardistance was normalized as a function of muscle size. Using thismeasure, both muscle size and internuclear distance werediminished in Dhc64C-depleted larvae compared with controls.However, internuclear distance was specifically reduced inclip190KG06490 and CLIP-190-depleted larvae, whereas muscle sizewas specifically reduced in lis1G10.14 (Fig. 8D-F). These datademonstrate a correlation between muscle output and both muscle

size and myonuclear position, suggesting that the defects seen inthe embryo are bona fide effects and not merely developmentaldelays.

DISCUSSIONWe have used the Drosophila musculature to investigate themechanisms that control muscle size and intracellular organization.We find that muscle length is regulated independently of thenumber of fusion events and demonstrate that perturbations thataffect embryonic muscle length correlate with decreased larvalmuscle size and poor muscle function. Additionally, we find thatintracellular organization, specifically myonuclear positioning, isessential for muscle function, consistent with our recentobservations (Metzger et al., 2012). Moreover, we have shown thatthe length and intracellular organization of the myofiber aremechanistically independent. Although a number of factors linkboth processes, we have identified factors that contribute solely tomuscle length or myonuclear position and demonstrate that we canindependently manipulate each feature.

Dynein regulates both muscle length and myonuclearpositioning. Some Dynein-interacting proteins, such as Dlc90F andPins, are necessary for both processes. That these factors regulateboth muscle length and myonuclear positioning suggests thatspecific aspects of muscle growth and the positioning of myonucleican indeed be coordinated. However, Lis1 affects muscle lengthspecifically, whereas CLIP-190 and Glued specifically affectmyonuclear position. Within the contexts of muscle length andmyonuclear position CLIP-190 and Lis1 do not genetically interact,illustrating that, although linked, the two processes aremechanistically distinct.

Our identification of CLIP-190 and Lis1 as regulators of Dyneinis not novel. CLIP-190 and Lis1 are known to interact with Dyneinboth physically and functionally, and they usually cooperate towarda single goal (Coquelle et al., 2002). Here, we provide the first

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Fig. 7. Microtubule organization is disrupted inCLIP-190 mutant embryos. (A)Confocal projectionsof a single hemisegment from stage 16 Drosophilaembryos immunostained for Tropomyosin (green) andTubulin (grayscale). The boxed regions are shown athigher magnification to the right. Yellow arrowsindicate individual and/or bundles of microtubules thatreach the muscle pole. Scale bars: 10m. (B)Thenumber of microtubules within 3m of the myofiberpole in the indicated genotypes. (C)The averageintensity of Tubulin immunofluorescence in the 3mnear the myotube pole in the indicated genotypes.Error bars represent s.d.; *P<0.05, compared withcontrol.

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example of CLIP-190 and Lis1 serving completely independent,Dynein-dependent functions.

Mechanistically, our data suggest that Dynein function, withrespect to the regulation of muscle length and myonuclearpositioning, is specified by Lis1 and CLIP-190 downstream of itslocalization. That Dynein localization to the myofiber pole isimportant is illustrated by pins (raps193) mutants in which Dyneindoes not accumulate at the myofiber pole, and we observe defectsin both muscle length and myonuclear positioning. This suggeststhat Pins recruits and stabilizes Dynein at the cortex as it doesduring mitosis (Gotta et al., 2003; Hughes et al., 2004). Lis1 alsoaffects Dynein localization, but in Lis1 mutants Dynein is localizedto the muscle pole and is tightly restricted to the pole comparedwith controls. We interpret this to mean that Lis1 is necessary for

retrograde trafficking of Dynein away from the myofiber pole. Wehypothesize that, in the absence of Dynein retrograde trafficking,cellular components accumulate at the extending muscle end, thusinhibiting the trafficking of factors necessary for further directedgrowth. Thus, when Dynein is incapable of moving cargo awayfrom the myofiber pole, muscles are shorter than in controlembryos. Indeed, a similar correlation between retrogradetrafficking and directed growth was recently reported duringmechanosensory bristle growth and axonal transport (Otani et al.,2011; Yi et al., 2011). Interestingly, it was recently reported thatdecreased retrograde transport of Dynein results in longer processesin both fibroblasts and neurons in culture (Rishal et al., 2012).

This contradictory finding raises several interesting questions. Isthere an opposing pathway in muscle or is another aspect of musclesize increased? For example, does the length of the muscle impactits volume? Complications due to muscle contraction in the liveembryo make such analysis difficult, however. Likewise, working invivo on a dynamically changing tissue located 30-150 m below thesurface of the developing embryo presents challenges for imagingthe rapid cellular processes that underlie motor activity, microtubuleorganization, organelle positioning and cell size. Nevertheless, thelink between different aspects of cell size and their relationships toeach other and to motor activity are being explored.

It is not clear why LT muscle growth/length is more sensitive toDynein activity than the VL muscles in the embryo. A simpleexplanation is that the maternally loaded Dynein persists longer inthe VL muscles than in the LT muscles. Additionally there mightbe a physical explanation. Although a cluster of potential tendoncells for the VL muscles exist, they are clustered at the segmentborder. Therefore, the size of the hemisegments, and thus thedistance from segment border to segment border, determinesmuscle length. Conversely, the cluster of potential tendon cells forthe LT muscles might be more broadly dispersed and the locationof the muscle pole at the time of tendon specification determinesthe length of the muscle. Under this hypothesis, inefficientextension of the LT muscles would result in the muscles beingshorter during tendon specification/maturation and therefore shorterthroughout embryonic development. Alternatively, differences inguidance/signaling systems could explain why the LT muscles, butnot the VL muscles, are shorter when Dynein activity iscompromised. Different signaling mechanisms are employed bythese different muscle types (Schnorrer and Dickson, 2004; Volk,1999; Schweitzer et al., 2010; Schejter and Baylies, 2010). Forexample, Derailed (Drl) plays a crucial role in the ability of LTmuscles to recognize their target (Callahan et al., 1996). Perhaps,altered trafficking of Drl or another factor in the signaling pathwaycauses slight, but significant, changes in LT muscle length.

It is interesting that the LT muscles are smaller in Dhc64C,Dlc90F, pins and Lis1 mutant embryos, but that other muscles areunaffected, whereas in larvae all muscles appear to be smaller.During the larval stages, muscles remain stably attached to tendoncells and grow through insulin signaling/Foxo- and dMyc-dependent pathways (Demontis and Perrimon, 2009). We interpretour observations to mean that Dynein and its regulatory proteinsDlc90F, Pins and Lis1 contribute to insulin receptor-mediatedmuscle growth as previously described (Huang et al., 2001; Varadiet al., 2003). Indeed, it would not be surprising to find that Dynein-dependent cellular trafficking is essential to that signaling pathway.

CLIP-190 does not dramatically affect Dynein localization, butit does affect microtubule organization, which, in turn, affectsnuclear positioning. CLIP-190 mutant embryos have fewermicrotubules at the myofiber pole, suggesting that, similar to its

3835RESEARCH ARTICLEDynein-dependent muscle formation

Fig. 8. Larval muscle organization and physiology are affected byDynein and associated proteins. (A,B)The average (A) andmaximum (B) speed of Drosophila larvae as they crawl towards astimulus. (C)Fluorescence images of VL3 muscles from L3 larvae thatwere used in locomotion assays just prior to dissection. White,phalloidin; green, Hoechst. Scale bar: 20m. (D)The average distancebetween nuclei in larval muscles from the indicated genotypes. (E)Thesurface area of the muscles in larvae of the indicated genotypes. (F)Thedistance between nuclei in the larval muscles normalized for musclesize. Error bars indicate s.d. *P<0.05, **P<0.01.

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functions in other systems, the role of CLIP-190 is to stabilizemicrotubule-cortex interactions (Neukirchen and Bradke, 2011;Watanabe et al., 2004), which Dynein then uses to move nucleitowards the myofiber pole. The specification of Dynein function,downstream of its localization, is novel. Although phospho-regulation has been shown to alter Dynein function during mitosis(Whyte et al., 2008) and the possibility for competitive regulationof Dynein has been suggested (McKenney et al., 2011), this is, toour knowledge, the first example in which Dynein at a singlelocation has its activity modified through interactions with uniquebinding partners. The ability to specify Dynein function withoutdramatically altering its localization is likely to be an importantfactor during development when temporal constraints are high.

With regards to physiology, the small, but significant, changesin myonuclear positioning and muscle size seen in the embryocontinue throughout larval development and are associated withimpaired muscle function. Additionally, that the same defects andimpairments are found in larvae that were depleted of Dynein, Lis1or CLIP-190 specifically in the muscle shows that the effects are,at least in part, muscle specific.

Many muscle myopathies are characterized by smaller myofibersand mispositioned nuclei. However, it is unclear whether thesepathologies are linked and which of these defects are paramount incausing the muscle weakness associated with these myopathies. Wehave shown that in Drosophila these two processes are linked viathe requirement for Dynein activity at the muscle pole. We havefurther shown that these two processes are mechanistically distinct,yet that both are necessary for muscle function. Together, these datasuggest that therapeutics aimed at improving the functionalcapacity of diseased muscles must counteract effects on bothmuscle size and myonuclear positioning. This highlightsDrosophila as an ideal model system with which to identify thegenes and mechanisms required for distinct aspects of musclemorphogenesis and to shed light on key features of muscle disease.

AcknowledgementsWe thank the Bloomington Stock Center and the Vienna Stock Center for flystocks and the Developmental Hybridoma Bank for antibodies.

FundingOur work is supported by Muscular Dystrophy Association (MDA) and NationalInstitutes of Health (NIH) [GM078318 to M.K.B.]. V.K.S. is supported by an NIHTraining Grant in Developmental Biology [T32HD060600]. Deposited in PMCfor release after 12 months.

Competing interests statementThe authors declare no competing financial interests.

Supplementary materialSupplementary material available online athttp://dev.biologists.org/lookup/suppl/doi:10.1242/dev.079178/-/DC1

ReferencesBate, M. (1990). The embryonic development of larval muscles in Drosophila.

Development 110, 791-804.Beckett, K. and Baylies, M. K. (2007). 3D analysis of founder cell and fusion

competent myoblast arrangements outlines a new model of myoblast fusion.Dev. Biol. 309, 113-125.

Bellen, H. J., Levis, R. W., Liao, G., He, Y., Carlson, J. W., Tsang, G., Evans-Holm, M., Hiesinger, P. R., Schulze, K. L., Rubin, G. M. et al. (2004). TheBDGP gene disruption project: single transposon insertions associated with 40%of Drosophila genes. Genetics 167, 761-781.

Brent, J. R., Werner, K. M. and McCabe, B. D. (2009). Drosophila larval NMJdissection. J. Vis. Exp. 24, 1107.

Caggese, C., Moschetti, R., Ragone, G., Barsanti, P. and Caizzi, R. (2001).dtctex-1, the Drosophila melanogaster homolog of a putative murine t-complexdistorter encoding a dynein light chain, is required for production of functionalsperm. Mol. Genet. Genomics 265, 436-444.

Callahan, C. A., Bonkovsky, J. L., Scully, A. L. and Thomas, J. B. (1996).derailed is required for muscle attachment site selection in Drosophila.Development 122, 2761-2777.

Coquelle, F. M., Caspi, M., Cordelières, F. P., Dompierre, J. P., Dujardin, D. L.,Koifman, C., Martin, P., Hoogenraad, C. C., Akhmanova, A., Galjart, N. etal. (2002). LIS1, CLIP-170’s key to the dynein/dynactin pathway. Mol. Cell. Biol.22, 3089-3102.

Demontis, F. and Perrimon, N. (2009). Integration of Insulin receptor/Foxosignaling and dMyc activity during muscle growth regulates body size inDrosophila. Development 36, 983-993.

Dietzl, G., Chen, D., Schnorrer, F., Su, K. C., Barinova, Y., Fellner, M., Gasser,B., Kinsey, K., Oppel, S., Scheiblauer, S. et al. (2007). A genome-widetransgenic RNAi library for conditional gene inactivation in Drosophila. Nature448, 151-156.

Etienne-Manneville, S. and Hall, A. (2001). Integrin-mediated activation ofCdc42 controls cell polarity in migrating astrocytes through PKCzeta. Cell 106,489-498.

Fitts, R. H., McDonald, K. S. and Schluter, J. M. (1991). The determinants ofskeletal muscle force and power: their adaptability with changes in activitypattern. J. Biomech. 24 Suppl. 1, 111-122.

Gepner, J., Li, M., Ludmann, S., Kortas, C., Boylan, K., Iyadurai, S. J.,McGrail, M. and Hays, T. S. (1996). Cytoplasmic dynein function is essential inDrosophila melanogaster. Genetics 142, 865-878.

Gill, S. R., Schroer, T. A., Szilak, I., Steuer, E. R., Sheetz, M. P. and Cleveland,D. W. (1991). Dynactin, a conserved, ubiquitously expressed component of anactivator of vesicle motility mediated by cytoplasmic dynein. J. Cell Biol. 115,1639-1650.

Goldstein, L. S. and Gunawardena, S. (2000). Flying through the drosophilacytoskeletal genome. J. Cell Biol. 150, 63F-68F.

Gönczy, P. (2002). Mechanisms of spindle positioning: focus on flies and worms.Trends Cell Biol. 12, 332-339.

Gotta, M., Dong, Y., Peterson, Y. K., Lanier, S. M. and Ahringer, J. (2003).Asymmetrically distributed C. elegans homologs of AGS3/PINS control spindleposition in the early embryo. Curr. Biol. 13, 1029-1037.

Grady, R. M., Starr, D. A., Ackerman, G. L., Sanes, J. R. and Han, M. (2005).Syne proteins anchor muscle nuclei at the neuromuscular junction. Proc. Natl.Acad. Sci. USA 102, 4359-4364.

Huang, J., Imamura, T. and Olefsky, J. M. (2001). Insulin can regulate GLUT4internalization by signaling to Rab5 and the motor protein dynein. Proc. Natl.Acad. Sci. USA 98, 13084-13089.

Hughes, J. R., Bullock, S. L. and Ish-Horowicz, D. (2004). Inscuteable mRNAlocalization is dynein-dependent and regulates apicobasal polarity and spindlelength in Drosophila neuroblasts. Curr. Biol. 14, 1950-1956.

Kraut, R. and Campos-Ortega, J. A. (1996). inscuteable, a neural precursor geneof Drosophila, encodes a candidate for a cytoskeleton adaptor protein. Dev. Biol.174, 65-81.

Lei, K., Zhang, X., Ding, X., Guo, X., Chen, M., Zhu, B., Xu, T., Zhuang, Y., Xu,R. and Han, M. (2009). SUN1 and SUN2 play critical but partially redundantroles in anchoring nuclei in skeletal muscle cells in mice. Proc. Natl. Acad. Sci.USA 106, 10207-10212.

Lei, Y. and Warrior, R. (2000). The Drosophila Lissencephaly1 (DLis1) gene isrequired for nuclear migration. Dev. Biol. 226, 57-72.

Liu, Z., Xie, T. and Steward, R. (1999). Lis1, the Drosophila homolog of a humanlissencephaly disease gene, is required for germline cell division and oocytedifferentiation. Development 126, 4477-4488.

Louis, M., Piccinotti, S. and Vosshall, L. B. (2008). High-resolution measurementof odor-driven behavior in Drosophila larvae. J. Vis. Exp. 3, 638.

McKenney, R. J., Vershinin, M., Kunwar, A., Vallee, R. B. and Gross, S. P.(2010). LIS1 and NudE induce a persistent dynein force-producing state. Cell141, 304-314.

McKenney, R. J., Weil, S. J., Scherer, J. and Vallee, R. B. (2011). Mutuallyexclusive cytoplasmic dynein regulation by NudE-Lis1 and dynactin. J. Biol.Chem. 286, 39615-39622.

Mesngon, M. T., Tarricone, C., Hebbar, S., Guillotte, A. M., Schmitt, E. W.,Lanier, L., Musacchio, A., King, S. J. and Smith, D. S. (2006). Regulation ofcytoplasmic dynein ATPase by Lis1. J. Neurosci. 26, 2132-2139.

Metzger, T., Gache, V., Xu, M., Cadot, B., Folker, E. S., Richardson, B. E.,Gomes, E. R. and Baylies, M. K. (2012). MAP and kinesin-dependent nuclearpositioning is required for skeletal muscle function. Nature 484, 120-124.

Neukirchen, D. and Bradke, F. (2011). Cytoplasmic linker proteins regulateneuronal polarization through microtubule and growth cone dynamics. J.Neurosci. 31, 1528-1538.

O’Rourke, S. M., Dorfman, M. D., Carter, J. C. and Bowerman, B. (2007).Dynein modifiers in C. elegans: light chains suppress conditional heavy chainmutants. PLoS Genet. 3, e128.

Otani, T., Oshima, K., Onishi, S., Takeda, M., Shinmyozu, K., Yonemura, S.and Hayashi, S. (2011). IKK regulates cell elongation through recyclingendosome shuttling. Dev. Cell 20, 219-232.

Palazzo, A. F., Joseph, H. L., Chen, Y. J., Dujardin, D. L., Alberts, A. S., Pfister,K. K., Vallee, R. B. and Gundersen, G. G. (2001). Cdc42, dynein, and dynactin

RESEARCH ARTICLE Development 139 (20)

DEVELO

PMENT

regulate MTOC reorientation independent of Rho-regulated microtubulestabilization. Curr. Biol. 11, 1536-1541.

Parmentier, M. L., Woods, D., Greig, S., Phan, P. G., Radovic, A., Bryant, P.and O’Kane, C. J. (2000). Rapsynoid/partner of inscuteable controls asymmetricdivision of larval neuroblasts in Drosophila. J. Neurosci. 20, RC84.

Pierson, C. R., Agrawal, P. B., Blasko, J. and Beggs, A. H. (2007). Myofiber sizecorrelates with MTM1 mutation type and outcome in X-linked myotubularmyopathy. Neuromuscul. Disord. 17, 562-568.

Prigozhina, N. L. and Waterman-Storer, C. M. (2004). Protein kinase D-mediated anterograde membrane trafficking is required for fibroblast motility.Curr. Biol. 14, 88-98.

Richardson, B. E., Beckett, K., Nowak, S. J. and Baylies, M. K. (2007).SCAR/WAVE and Arp2/3 are crucial for cytoskeletal remodeling at the site ofmyoblast fusion. Development 134, 4357-4367.

Rishal, I., Kam, N., Perry, R. B., Shinder, V., Fisher, E. M., Schiavo, G. andFainzilber, M. (2012). A motor driven mechanism for cell length sensing. CellRep. 1, 608-616.

Romero, N. B. (2010). Centronuclear myopathies: a widening concept.Neuromuscul. Disord. 20, 223-228.

Rommel, C., Bodine, S. C., Clarke, B. A., Rossman, R., Nunez, L., Stitt, T. N.,Yancopoulos, G. D. and Glass, D. J. (2001). Mediation of IGF-1-inducedskeletal myotube hypertrophy by PI(3)K/Akt/mTOR and PI(3)K/Akt/GSK3pathways. Nat. Cell Biol. 3, 1009-1013.

Schejter, E. D. and Baylies, M. K. (2010). Born to run: creating the muscle fiber.Curr. Opin. Cell Biol. 22, 566-574.

Schmoranzer, J., Kreitzer, G. and Simon, S. M. (2003). Migrating fibroblastsperform polarized, microtubule-dependent exocytosis towards the leading edge.J. Cell Sci. 116, 4513-4519.

Schnorrer, F. and Dickson, B. J. (2004). Muscle building; mechanisms of myotubeguidance and attachment site selection. Dev. Cell 7, 9-20.

Schweitzer, R., Zelzer, E. and Volk, T. (2010). Connecting muscles to tendons:tendons and musculoskeletal development in flies and vertebrates. Development137, 2807-2817.

Sheeman, B., Carvalho, P., Sagot, I., Geiser, J., Kho, D., Hoyt, M. A. andPellman, D. (2003). Determinants of S. cerevisiae dynein localization andactivation: implications for the mechanism of spindle positioning. Curr. Biol. 13,364-372.

Sönnichsen, B., Koski, L. B., Walsh, A., Marschall, P., Neumann, B., Brehm,M., Alleaume, A. M., Artelt, J., Bettencourt, P., Cassin, E. et al. (2005). Full-genome RNAi profiling of early embryogenesis in Caenorhabditis elegans.Nature 434, 462-469.

Squire, J. M. (1997). Architecture and function in the muscle sarcomere. Curr.Opin. Struct. Biol. 7, 247-257.

Tanenbaum, M. E., Macurek, L., Galjart, N. and Medema, R. H. (2008).Dynein, Lis1 and CLIP-170 counteract Eg5-dependent centrosome separationduring bipolar spindle assembly. EMBO J. 27, 3235-3245.

Tsai, J. W., Bremner, K. H. and Vallee, R. B. (2007). Dual subcellular roles forLIS1 and dynein in radial neuronal migration in live brain tissue. Nat. Neurosci.10, 970-979.

Varadi, A., Tsuboi, T., Johnson-Cadwell, L. I., Allan, V. J. and Rutter, G. A.(2003). Kinesin I and cytoplasmic dynein orchestrate glucose-stimulated insulin-containing vesicle movements in clonal MIN6 beta-cells. Biochem. Biophys. Res.Commun. 311, 272-282.

Vaughan, P. S., Miura, P., Henderson, M., Byrne, B. and Vaughan, K. T.(2002). A role for regulated binding of p150(Glued) to microtubule plus ends inorganelle transport. J. Cell Biol. 158, 305-319.

Volk, T. (1999). Singling out Drosophila tendon cells: a dialogue between twodistinct cell types. Trends Genet. 15, 448-453.

Watanabe, T., Wang, S., Noritake, J., Sato, K., Fukata, M., Takefuji, M.,Nakagawa, M., Izumi, N., Akiyama, T. and Kaibuchi, K. (2004). Interactionwith IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cellpolarization and migration. Dev. Cell 7, 871-883.

Waterman-Storer, C. M., Karki, S. B., Kuznetsov, S. A., Tabb, J. S., Weiss, D.G., Langford, G. M. and Holzbaur, E. L. (1997). The interaction betweencytoplasmic dynein and dynactin is required for fast axonal transport. Proc. Natl.Acad. Sci. USA 94, 12180-12185.

Whyte, J., Bader, J. R., Tauhata, S. B., Raycroft, M., Hornick, J., Pfister, K. K.,Lane, W. S., Chan, G. K., Hinchcliffe, E. H., Vaughan, P. S. et al. (2008).Phosphorylation regulates targeting of cytoplasmic dynein to kinetochoresduring mitosis. J. Cell Biol. 183, 819-834.

Yi, J. Y., Ori-McKenney, K. M., McKenney, R. J., Vershinin, M., Gross, S. P.and Vallee, R. B. (2011). High-resolution imaging reveals indirect coordinationof opposite motors and a role for LIS1 in high-load axonal transport. J. Cell Biol.195, 193-201.

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