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VCP-dependent muscle degeneration is linked to defects in a dynamic tubular lysosomal network in vivo

  1. Alyssa E Johnson
  2. Huidy Shu
  3. Anna G Hauswirth
  4. Amy Tong
  5. Graeme W Davis  Is a corresponding author
  1. University of California, San Francisco, United States
Research Article
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Cite this article as: eLife 2015;4:e07366 doi: 10.7554/eLife.07366

Abstract

Lysosomes are classically viewed as vesicular structures to which cargos are delivered for degradation. Here, we identify a network of dynamic, tubular lysosomes that extends throughout Drosophila muscle, in vivo. Live imaging reveals that autophagosomes merge with tubular lysosomes and that lysosomal membranes undergo extension, retraction, fusion and fission. The dynamics and integrity of this tubular lysosomal network requires VCP, an AAA-ATPase that, when mutated, causes degenerative diseases of muscle, bone and neurons. We show that human VCP rescues the defects caused by loss of Drosophila VCP and overexpression of disease relevant VCP transgenes dismantles tubular lysosomes, linking tubular lysosome dysfunction to human VCP-related diseases. Finally, disruption of tubular lysosomes correlates with impaired autophagosome-lysosome fusion, increased cytoplasmic poly-ubiquitin aggregates, lipofuscin material, damaged mitochondria and impaired muscle function. We propose that VCP sustains sarcoplasmic proteostasis, in part, by controlling the integrity of a dynamic tubular lysosomal network.

https://doi.org/10.7554/eLife.07366.001

eLife digest

Mutations in a gene that produces a protein called Valosin-containing protein (VCP for short) causes degenerative diseases that affect the brain, muscle and bone. In nearly half of the individuals with these VCP-related diseases—which can also result in dementia, Paget's disease of the bone and amyotrophic lateral sclerosis (ALS)—the first symptom is muscle weakness. Currently, very little is known about how VCP affects muscles.

Patients with VCP-related diseases often have problems clearing damaged proteins from their cells, and recent research suggests that VCP is important for forming a cellular structure known as a lysosome. Lysosomes contain powerful enzymes that destroy damaged proteins and other cellular structures that would otherwise accumulate in the cells. In most cells, lysosomes look like bubble-like compartments called vesicles. However, in some types of cells lysosomes have been observed to form a network of tubules that extend throughout the cell interior. However, it remains unclear what these tubules do, how they form in cells and whether they are altered in disease.

Johnson et al. analyzed lysosomes in the muscle of the fruit fly species Drosophila melanogaster and discovered that lysosomes were in the form of a network of tubules that spread throughout each muscle cell. These tubules constantly changed in living muscles; extending, retracting, breaking and merging to form a large tubular lysosome network. When Johnson et al. reduced the amount of VCP produced by the muscle cells, via a method called RNA interference, the lysosome tubules broke down into vesicles that were no longer constantly changing. Modifying these defective fly muscle cells so that they produced the human VCP protein caused the tubules to form again. These results suggest that the human and fly VCP proteins are very similar and that they play a key role in either the ability of lysosomes to form tubules or the maintenance of existing tubules.

Johnson et al. then engineered flies to produce a version of the VCP protein that had mutations commonly seen in individuals with degenerative diseases. Lysosome tubules did not form correctly in the muscle cells of these flies. These flies also had other abnormalities; for example, their cells showed a great build-up of damaged proteins, and their ability to move their muscles was weaker.

These findings suggest that a network of lysosomal tubules is necessary for healthy muscle cells, but how and why these tubular networks are formed or maintained is still mysterious. What causes lysosomal membranes to form tubules? How do they break and fuse? And why are they necessary? Genetic experiments in fruit flies will be a great place to discover these mechanisms and understand the links to degenerative diseases in humans.

https://doi.org/10.7554/eLife.07366.002

Introduction

Valosin-containing protein (VCP), the homologue of yeast Cdc48, is the causative gene for a multisystem degenerative disease that was originally termed IBMPFD to encompass the wide range of debilitating clinical outcomes, including inclusion body myopathy (IBM), Paget's disease of the bone (PDB) and frontotemporal dementia (FD) (Watts et al., 2004). Recently, the list of degenerative disorders that are associated with VCP mutations has expanded to include amyotrophic lateral sclerosis (ALS) (Abramzon et al., 2012), spastic paraplegia (Clemen et al., 2010), scapuloperoneal muscular dystrophy (Liewluck et al., 2014) and Charcot-Marie-Tooth disease (Gonzalez et al., 2014). Currently, there are no viable treatments available to slow or halt progression of VCP-related diseases.

Muscle weakness is the first presenting symptom in over 50% of VCP disease patients (Weihl et al., 2009), yet very little is known about the muscle pathology of VCP-related diseases. Muscle biopsies from patients with VCP-related diseases display an accumulation of cytoplasmic poly-ubiquitin aggregates (Watts et al., 2004; Weihl et al., 2009; Dolan et al., 2011), suggesting a major defect in protein clearance. VCP is an AAA-ATPase that has essential functions in ubiquitin-dependent proteolysis. But, pathogenic mutations in VCP do not seem to impair the UPS or ERAD protein degradation pathways (Tresse et al., 2010a; Chang et al., 2011). More recently, VCP has been implicated in autophagy. Specifically, over-expression of VCP mutant transgenes with disease causing mutations leads to an accumulation of autophagosomes (Ju et al., 2009; Tresse et al., 2010a), suggesting that VCP functions in processes related to the maturation or fusion of autophagosomes with lysosomes.

Lysosomes are the major cellular degradation sites for clearing damaged proteins and organelles. Lysosomes are classically thought to be vesicular organelles, where they serve as depots for cargo delivered via endosomes or autophagosomes. However, in certain systems lysosomes have been observed to adopt non-vesicular morphologies. A particularly dramatic example has been observed in a subset of bone-derived cultured macrophages, where lysosomes form abundant, extended tubules that radiate from the cell center and, in some cases, form an interconnected web throughout the cytoplasm (Swanson et al., 1987a; Knapp and Swanson, 1990). Additionally, there are examples where cellular stress, particularly the induction of high levels of autophagy, induces lysosomal membranes to tubulate and undergo scission to produce new vesicular lysosomes, a process referred to as autophagic lysosome reformation (ALR) (Yu et al., 2010). Despite these observations, lysosome tubules have received little attention and it remains unclear to what extent lysosome tubulation occurs in different cell types and what purpose it serves in vivo. Moreover, the molecular repertoire of factors required for lysosome tubule formation is virtually unknown.

Here, we employ fluorescently tagged lysosomal and autophagic markers to study the autophagy-lysosome system in Drosophila muscle cells and investigate the muscle pathology of VCP-related diseases. Remarkably, we find that lysosomes adopt a dynamic, tubular morphology that ramifies throughout the entire sarcoplasm of Drosophila muscle, in vivo. We find that VCP is required for the integrity and dynamics of this tubular network. Disruption of lysosome tubules correlates with defects in autophagosome-lysosome fusion, increased poly-ubiquitin aggregates and the accumulation of lipofuscin material in the sarcoplasm. We show that the human VCP homolog can rescue lysosomal tubulation following loss of VCP in Drosophila muscle, indicating that the functions of VCP in lysosome tubulation are conserved. Finally, we demonstrate that homologous mutations that cause VCP diseases in human patients disrupt the lysosome tubular lattice, suggesting that disruption of lysosome tubules contributes to VCP mutant pathogenesis. Taken together, our data establish a functional link between lysosome tubule dysfunction and the pathology of VCP-related degenerative diseases.

Results

Drosophila sarcoplasmic lysosomes form an extended dynamic tubular array in vivo

To visualize muscle lysosomes in vivo, we expressed RFP-tagged Spinster, which has previously been defined as a late endosomal/lysosomal transmembrane protein (Sweeney and Davis, 2002; Dermaut et al., 2005). Remarkably, when Spinster-RFP is expressed in Drosophila muscle, Spin-RFP localizes to an expansive tubular network (Figure 1A–C and Video 1). Tubules were found evenly distributed throughout the sarcoplasm and formed a web of connections with other tubules (Figure 1B,C). Tubules were observed in every muscle and there were no apparent differences in the tubule abundance or architecture between different muscles. We also observed enlarged vesicular compartments at tubule intersections throughout the muscle (Figure 1C). This network is highly sensitive to all chemical fixation conditions that we have attempted. When subjected to fixation, the tubule lattice collapses (Figure 1D), leaving behind distributed round, Spinster-positive compartments that resemble classically defined late-endosomal, lysosomal structures (Sweeney and Davis, 2002; Dermaut et al., 2005). Thus, live imaging is essential to study the function and relevance of this Spin-positive, tubular network.

Lysosomes adopt an extended dynamic tubular array in Drosophila sarcoplasms.

(A) Muscles of third instar larvae from segment A2. (B) Representative live image of Spin-RFP expressed in muscles at 63× magnification. Muscle 4 (B) is shown. DNA was stained with Hoescht. (C) Representative live image of Spin-RFP expressed in muscles using the muscle-specific MHC-Gal4 driver. DNA was stained with Hoescht. (D) Representative image of Spin-GFP localization in a muscle that was fixed with 4% PFA prior to imaging. (E) Time-lapse images of the Spin-RFP network. Time 0 is represented in magenta, and the 5 min time-point is represented in green. The 2 time points were merged to show new and lost tubule formations over the course of 5 min. Arrows indicate examples of de novo tubule formations and the asterisk indicates a retracted tubule. (F) Representative time-lapse sequence of a Spin-GFP tubule fission event. (G) Representative time-lapse sequence of a Spin-GFP tubule fusion event. In the last frame, a de novo tubule can be seen extruding from the middle of a pre-existing tubule. (HJ) Spin-RFP localization in muscles treated with DMSO (H), Nocodazole (I) or LatA (J). (K) Spin-RFP localization in muscles expressing Clathrin heavy chain (Chc) RNAi.

https://doi.org/10.7554/eLife.07366.003
Video 1
Spin-RFP tubular network in Drosophila muscle.

Spin-RFP was expressed in muscles and imaged live. Average Z-stacks were assembled to produce a 3D volume projection and various angles of the projections are shown.

https://doi.org/10.7554/eLife.07366.004

We next explored the dynamics of this tubular network in time-lapse videos. We find evidence of tubule extension, retraction, scission and fusion (Figure 1E–G). The dominant activity in the network is the dynamic extension and retraction of individual tubules throughout the sarcoplasm (Figure 1E and Video 2). Often the same tubule was observed to extend and retract, while surrounding tubules remained constant. The impression is that the lattice has both stable, interconnected tubules and dynamic elements that create new connections or retract once a connection is broken. In less frequent instances, we observed scission events from the end of a tubule, resulting in a mobile vesicle (Figure 1F). In a corollary phenomenon, we observed tubule extensions that resulted in tubules fusing to larger nodes in the network (Figure 1G). Finally, we observed de novo formation of tubules extruding from the side of existing tubules (Figure 1G, last panel) demonstrating that tubulation can originate from existing tubules, not just pre-existing nodes. Importantly, the existence of a tubular network and tubule dynamics are not an artifact of a dissected neuromuscular preparation. We observed an identical, dynamic tubular lysosomal network in intact larvae that were imaged through the cuticle while restrained in a microfluidics chamber (Video 3).

Video 2
Spin-RFP tubule dynamics in Drosophila muscle.

Representative time-lapse video of Spin-RFP expressed in muscles. Frames were taken at 10 s intervals.

https://doi.org/10.7554/eLife.07366.005
Video 3
Spin-RFP tubule dynamics in an intact larva.

A whole un-dissected larva was immobilized in a mircrofluidics chamber and Spin-RFP was imaged in the body-wall muscle through the transparent cuticle. Frames were taken at 10 s intervals.

https://doi.org/10.7554/eLife.07366.006

The observed lysosomal network structure and dynamics are consistent with the involvement of either the actin or microtubule cytoskeletons. First, we tested whether the tubules require an intact microtubule cytoskeleton. Treatment with nocodazole for 1 hr completely abolished tubules, indicating that the tubules require microtubule support (Figure 1H,I). In contrast, disruption of the actin cytoskeleton with latrunculin A did not have a significant effect on the tubular network, indicating a non-essential role for the actin cytoskeleton in maintaining lysosome tubules (Figure 1J). Finally, we tested whether Clathrin is essential for tubular network integrity. Clathrin has the capacity to shape membranes and was recently implicated in the process of ALR in cultured mammalian cells (Rong et al., 2012), a process that involves limited tubulation from auto-lysosomal compartments. Expression of Clathrin heavy chain RNAi (Chc-RNAi) completely disrupted network integrity (Figure 1K).

Although Spinster was characterized previously as a lysosomal marker, the tubular structures labeled by Spinster are dramatically different from the classical view of vesicular lysosomes. Therefore, we performed additional experiments to validate that the tubular network is lysosomal. First, we co-imaged Spin-GFP with the low pH fluorescent probe Lysotracker and found complete co-localization (Figure 2A), indicating that the Spinster network is acidic. To verify that the observed tubular network is not an artifact of Spin-RFP over-expression, we stained wild type muscles with Lysotracker to examine lysosome morphology under wild type conditions. Lysotracker staining confirmed the existence of this tubular network in wild type muscle (Figure 2B). We also note that the tubule intersections stained more intensely for Lysotracker than the tubules themselves. This could be due to increased tubule volume at the intersections, or these sites might actually have a lower pH than the tubules themselves. Finally, we co-imaged Spin-RFP with several other organelle markers to verify that Spinster specifically labels lysosomes and does not co-stain other organelles. ER and mitochondria also form tubule structures, but when we co-imaged Spin-RFP with ER-tracker and Mito-tracker fluorescent dyes, Spin-RFP tubules did not co-localize with either ER or mitochondria tubules (Figure 2C,D). Instead, Spin-RFP tubules were interwoven between mitochondria and ER tubules. Additionally, Spin-RFP did not co-localize with markers for early endosomes (YFP-Rab5), recycling endosomes (Rab11-GFP), medial Golgi (ManII-GFP) or trans Golgi (GalT-YFP) (Figure 2E–H). We note that Golgi organization in muscles forms vesicular structures rather than the classical Golgi stacks that are observed in most cell types and this is consistent with Golgi organization that has been observed in vertebrate skeletal muscles (Ralston et al., 2001). Collectively, we have identified and characterized a dynamic tubular lysosomal network that permeates the entire sarcoplasm of Drosophila body-wall muscle in vivo.

Spin-RFP tubules do not co-localize with mitochondria, ER, golgi or early endosomes.

(A) Co-imaging of Spin-GFP and Lysotracker Red staining. (B) Lysotracker staining of wild type muscles. (CH) Co-imaging of Spin-RFP with ER tracker (C), Mito tracker (D), YFP-Rab5 (E), Rab11-GFP (F), ManII-GFP (G), GalT-YFP (H).

https://doi.org/10.7554/eLife.07366.007

VCP is required for the integrity and dynamics of the extended tubular lysosomal network

To investigate the molecular underpinnings of the observed lysosomal tubule dynamics, we pursued a candidate-based RNAi screen to identify genes required for lysosome tubulation. We focused on genes that have been implicated in the autophagy-lysosome system and identified the AAA-ATPase VCP as being required for the integrity of the entire tubular lysosome network. Specifically, inhibiting VCP expression by RNAi abolished lysosome tubules, leaving behind vesicles throughout the sarcoplasm (Figure 3A,B). The lysosome vesicles were irregular in their size and shape and appeared clustered, rather than uniformly distributed throughout the sarcoplasm. To determine whether the catalytic ATPase function of VCP is required for the tubular network integrity, we employed a VCP-selective inhibitor DBeQ (Chou et al., 2011). Acute inhibition of VCP with DBeQ completely disrupted the tubular network after 3 hr (Figure 3C,D), demonstrating the required catalytic function of VCP. Furthermore, time-lapse imaging of Spin-GFP after treatment with DBeQ for 4 hr revealed that the remaining vesicular lysosome structures are completely static (Video 4).

VCP inhibition disrupts the lysosome tubule lattice and human VCP rescues this defect.

(A) Representative live image of Spin-RFP expressed in muscle using the muscle-specific BG57-Gal4 driver. (B) Live image of Spin-RFP in muscles expressing VCP-RNAi using the muscle-specific BG57-Gal4 driver. (C, D) Live images of Spin-RFP expressed in muscles that were treated with DMSO (C) or the VCP-specific inhibitor DBeQ (D) for 4 hr. (E) Live image of Spin-GFP expressed in muscles using the muscle-specific BG57-Gal4 driver. (F) Live image of Spin-GFP in muscles expressing VCP-RNAi using the muscle-specific BG57-Gal4 driver. (G) Live image of Spin-GFP in muscles that co-express VCP-RNAi and human VCP (hVCP) using the muscle-specific BG57-Gal4 driver. (HJ) Lysotracker staining in wild type (H) muscles or muscles expressing parkin-RNAi (I) or tbph-RNAi (J). (K, L) Spin-RFP localization in muscles treated with DMSO (K) or tunicamycin (TM) (L). (M) Western blot analysis of total Hsc70/BiP protein levels. Tubulin serves as a loading control.

https://doi.org/10.7554/eLife.07366.008
Video 4
Spin-GFP dynamics in muscles expressing VCP-RNAi.

Representative time-lapse video of Spin-GFP in muscles expressing VCP-RNAi. Frames were taken at 10 s intervals.

https://doi.org/10.7554/eLife.07366.009

To determine if the role of VCP in maintaining the tubular lysosomal network in muscle cells is conserved, we overexpressed human VCP in dVCPRNAi muscles. Human VCP should be completely resistant to dVCPRNAi due to lack of extended stretches of nucleotide identity. Over-expressing human VCP in dVCPRNAi muscles rescued the formation of lysosome tubules in every muscle (Figure 3E–G). These rescue data confirm that VCP knockdown is the cause of disrupted lysosomal network integrity and demonstrate that VCP-dependent activity is conserved in the human VCP protein.

Since VCP knockdown destroys the sarcoplasmic tubular lysosomal network prior to obvious cellular degeneration, we speculated that dismantling of the network could be a precursor to cellular degeneration. However, it is also possible that loss of this network is a secondary correlate of impaired muscle health. To address this issue, we examined two additional RNAi-mediated conditions that cause muscle degeneration, looking for the presence or absence of tubular lysosomes. Specifically, we expressed RNAi against parkin and tbph, Drosophila orthologues of genes linked to Parkinson's and ALS, respectively. These RNAi have been shown to cause muscle degeneration in Drosophila (Diaper et al., 2013; Cornelissen et al., 2014). Remarkably, we did not observe any significant effect on lysosome tubules (Figure 3H–J) when parkin-RNAi and tbph-RNAi are expressed with BG57-Gal4, the same Gal4 line used to express VCP-RNAi throughout our studies.

VCP has well-established roles in the ERAD pathway and loss of VCP causes ER stress (Wójcik et al., 2006). Thus, it is possible that loss of tubular lysosomes is caused indirectly by ER stress. To address this possibility, we treated muscles with tunicamycin (TM) to induce ER stress by another means and examined the tubular network. Upon treatment with TM for 4 hr, the Spin-RFP tubular network remained intact (Figure 3K,L). We verified that ER stress was induced by examining levels of Hsc70/BiP, a marker of ER stress, in protein lysates derived from the same animals that were used for imaging. Total protein levels of Hsc70/BiP were significantly increased in both TM and DBeQ treated muscles (Figure 3M). Thus, tubular network disruption is not a byproduct of ER stress. Taken together, our data are consistent with three conclusions: (1) VCP loss of function disrupts the integrity of the tubular lysosomal network, (2) the role of VCP in maintaining tubular lysosomes is conserved and (3) disruption of the tubular network is specific for VCP loss of function and not a secondary byproduct of muscle degeneration or ER stress.

Autophagosome membranes co-localize with the tubular lysosome network

Muscle biopsies from patients with VCP-related disease display an accumulation of cytoplasmic poly-ubiquitin aggregates (Watts et al., 2004; Weihl et al., 2009; Dolan et al., 2011), suggesting a defect in protein clearance. This led us to explore the intersection of the observed tubular lysosomal network and autophagy. First, we co-imaged Spin-GFP with the autophagosome membrane marker mCherry-Atg8a/LC3 to determine the relationship between autophagosomes and the tubular lysosome network. Remarkably, we find that mCherry-Atg8a precisely co-localizes with Spin-GFP-labeled tubules (Figure 4A). The mCherry-Atg8a labeling is widely distributed within the network, but does not label the full extent of every tubule (Figure 4A). Time-lapse imaging revealed mCherry-Atg8a labeling is dynamic within Spin-GFP tubules (Video 5). Some mCherry-Atg8a puncta appear to traffic through the Spin-GFP tubules suggesting that autophagosome membranes and/or cargos are dynamic within the tubular lysosome network (Figure 4B). Atg8 localizes to the autophagophore membrane prior to the formation of an enclosed autophagosome, at which point Atg8 is shed from the outer autophagosome membrane (Xie and Klionsky, 2007). After autophagosomes fuse with lysosomes, lysosomal enzymes degrade the inner autophagosome membrane containing Atg8 (Xie and Klionsky, 2007). Given this sequence of events, we propose that the mCherry-Atg8a positive regions of the lysosome tubules represent ongoing degradation of autophagosome membranes following fusion with the Spin-GFP positive lysosomal network.

Autophagosomes co-localize with the tubular lysosomal network.

(A) Representative live image of Spin-GFP and mCherry-Atg8a co-expressed in muscles using the muscle-specific BG57-Gal4 driver. (B) Representative time-lapse sequence of Spin-GFP and mCherry-Atg8a in muscle. The arrow follows a mCherry-Atg8a positive puncta trafficking along a Spin-GFP tubule. (C) Spin-GFP and mCherry-Atg8a no longer co-localize in muscles expressing VCP-RNAi. White box indicates region shown at higher magnification and separate channels at right. D. Live image of GFP-mCherry-Atg8a in muscles treated with DMSO for 3 hr. Note the lack of GFP signal. (E) Live image of GFP-mCherry-Atg8a in muscles treated with the V-ATPase specific inhibitor Concanamycin A (ConA) for 3 hr. Note the presence of GFP-positive tubules. (F) Live image of GFP-mCherry-Atg8a in muscles treated with the VCP-specific inhibitor DBeQ for 3 hr. Note the presence of GFP-positive vesicles. (G) Spin-RFP localization in WT muscles or muscles expressing Atg7-RNAi using the muscle specific BG57-Gal4 driver.

https://doi.org/10.7554/eLife.07366.010
Video 5
Spin-GFP and mCherry-Atg8a dynamics in Drosophila muscle.

Representative time-lapse video of Spin-GFP and mCherry-Atg8 co-expressed in muscles. Frames were taken at 10 s intervals.

https://doi.org/10.7554/eLife.07366.011

Next, we explored the consequence of disrupting VCP activity on Atg8 and Spin-GFP co-localization. When VCP was knocked down and the tubular-lysosomal network was eliminated, Atg8 no longer co-localized with Spin-GFP (Figure 4C). Rather, Atg8-positive vesicles were found closely apposed to Spin-positive, vesicular lysosomal compartments. This close apposition suggests that the autophagosomes can identify and potentially dock against the lysosomal membranes, but fusion of the autophagosome with the lysosome is defective. This is consistent with an established role for the yeast VCP homolog, Cdc48, in membrane fusion (Latterich et al., 1995).

To test whether tubules that are positive for both mCherry-Atg8a and Spin-GFP are, indeed, a consequence of fused auto-lysosomes, we imaged a dual fluorescent reporter GFP-mCherry-Atg8a (Nezis et al., 2010). In neutral pH conditions, both GFP and mCherry fluoresce. However, in acidic environments such as in the lysosomal lumen, GFP fluorescence is quenched and only mCherry is observed. Thus, autophagosomes that are fused with lysosomes will exhibit mCherry but not GFP fluorescence. It is important to note that Spin-GFP fluorescence is not quenched because the C-terminal GFP tag resides on the cytoplasmic face of lysosomes (Dermaut et al., 2005). When we expressed the dual fluorescence reporter in muscles we found no detectable GFP fluorescence in the tubules that label strongly for mCherry-Atg8a (Figure 4D), consistent with other data indicating that the tubules are acidic. To further test the acidic nature of the tubules, we treated muscles with concanamycin A, a specific inhibitor of the lysosomal V-ATPase that is required for lysosome acidification (Huss et al., 2002). Upon treatment with ConA for 3 hr, we observed GFP and mCherry positive tubules (Figure 4E). Together, these data further verify that the lysosome tubules are acidic and also demonstrate that acidification is not necessary to maintain the structure of the tubules.

Again, we explored the consequence of disrupting VCP activity. Inhibiting VCP function with DBeQ created GFP-positive vesicles (Figure 4F) that must be non-acidified compartments, a finding that is consistent with the existence of autophagosomes that have not yet fused with Spin-positive lysosomes. Since the appearance of GFP-positive vesicles occurs in a time frame of minutes to hours, it suggests that there is a continual flux of material through the auto-lysosomal system in muscle cells at steady state.

Finally, we examined whether autophagy influx was required to induce or maintain lysosome tubules. Expression of Atg7-RNAi, an RNAi line that has been shown previously to inhibit autophagosome assembly (Ren et al., 2009), did not affect lysosome tubules (Figure 4G). Thus, autophagy induction is not a prerequisite for lysosome tubules. Taken together, these data confirm previous reports that VCP is required for autophagosome-lysosome fusion (Tresse et al., 2010b) and further suggest that VCP loss abrogates autophagosome-lysosome fusion by affecting the structural properties of lysosomes.

VCP localizes to auto-lysosomes

Previous studies have reported that, when over-expressed in cultured cells, mammalian VCP localizes diffusely at the nucleus and throughout the cytoplasm (Vesa et al., 2009; Tresse et al., 2010a; Wang et al., 2011). To examine VCP localization in live muscles, we generated a UAS-VCP-Venus transgenic fly. Similar to previous reports for mammalian VCP, we observed abundant VCP localization in and around the nucleus and diffusely in the cytoplasm (Figure 5—figure supplement 1). But, VCP-Venus also concentrated at structures that are labeled by either Spin-RFP (Figure 5A) or mCherry-Atg8a (Figure 5B), demonstrating that VCP localizes to auto-lysosomes.

Figure 5 with 1 supplement see all
VCP co-localizes with the tubular auto-lysosomes.

(A) Representative live image of VCP-Venus and Spin-RFP expressed in muscles using the muscle-specific BG57-Gal4 driver. White box indicates region shown at higher magnification and separate channels at right. (B) Representative live image of VCP-Venus and mCherry-Atg8a expressed in muscles using the muscle-specific BG57-Gal4 driver. Inset as in A. (C, D) VCP-Venus and mCherry-Atg8a localization in muscles treated with the VCP inhibitor DBeQ (C) or the proteasome inhibitor MG132 (D) for 3 hr. (E) Western blot analysis of total VCP protein levels from muscles in the treatments indicated. Tubulin serves as a loading control. (F) Representative time-lapse sequence of VCP-Venus and mCherry-Atg8a after MG132 was washed out. The arrow indicates a tubule extending from a mCherry-Atg8a positive vesicle.

https://doi.org/10.7554/eLife.07366.012

Because inhibiting the catalytic function of VCP leads to tubule deterioration, we tested whether inhibiting the catalytic function of VCP would affect its localization to auto-lysosomes. Surprisingly, inhibiting VCP activity with DBeQ for 2 hr triggered the formation of rod-shaped VCP aggregates that effectively sequester VCP-Venus from the sarcoplasm (Figure 5C). The formation of these aggregates is striking, as they resemble rod-like structures characteristic of prion aggregates. We considered two possible reasons that application of DBeQ might increase the propensity for VCP to aggregate. First, DBeQ binding to VCP might initiate aggregation directly by altering the solubility of VCP. Alternatively, the aggregation could be caused by the functions of VCP in other contexts, such as proteasome-dependent protein degradation. Remarkably, when we applied the proteasome inhibitor MG132, VCP-Venus rapidly aggregated in a manner identical to that observed following DBeQ incubation (Figure 5D). VCP aggregation was not due to increased VCP protein levels as a result of proteasome inhibition, because total VCP protein did not increase significantly upon MG132 or DBeQ treatment (Figure 5E).

While the significance of VCP-Venus aggregates remains uncertain, this phenotype provided us with a means to rapidly and reversibly sequester VCP protein and control its access to auto-lysosomal membranes. Since VCP exists as a hexamer, MG132 incubation in muscles over-expressing VCP-Venus should sequester both wild type and Venus-tagged VCP. Application of MG132 induced VCP-Venus aggregate formation, which correlated with dissolution of the tubular lysosomal network (Figure 5D). When MG132 was washed out, VCP aggregates dissolved within 20 min and, as they disappeared, cytoplasmic VCP fluorescence intensity increased (Figure 5F and Video 6). During this time, cytoplasmic VCP-Venus accumulated at autophagosomes/lysosomes (mCherry-Atg8a) and tubules began to reform (Figure 5F and Video 6). These data indicate that VCP localizes to auto-lysosomes, where it could participate in auto-lysosomes tubulation.

Video 6
VCP-Venus and mCherry-Atg8a dynamics after MG132 wash out.

Muscles co-expressing VCP-Venus and mCherry-Atg8 were treated with the proteasome inhibitor MG132 for 3 hr. MG132 was washed out and time-lapse images were taken every 10 s.

https://doi.org/10.7554/eLife.07366.014

Disrupted lysosome tubules correlate with muscle weakness and autophagy/lysosome defects

We next investigated the consequence of disrupting the auto-lysosome tubule network. We first examined overall muscle function. When VCP RNAi was expressed specifically in muscle, muscle wasting was apparent and third instar larvae exhibited a severe impairment in their ability to move (Figure 6—figure supplement 1A,B). When prodded, the animals would move, but their movements were slow and only lasted for short periods of time. The defect in their motility is not due to defects in the nervous system because synaptic transmission at the NMJ remained intact (Figure 6—figure supplement 1C,D). These data are consistent with compromised muscle function that parallels the muscle weakness observed in human VCP-related diseases.

Next, we investigated the degradation capacity of the collapsed tubular lysosomes. For lysosomes to degrade their cargo they must be acidified and the proteolytic enzymes must be present. We first examined the acidification of the lysosomes by co-imaging Spin-GFP with Lysotracker in VCP-RNAi animals and found that the enlarged Spin-GFP vesicles also co-stained with Lysotracker (Figure 6—figure supplement 2A), indicating that they are acidic. Then, we examined whether lysosome enzymes were delivered properly to the lysosomes. Normally, the lysosomal enzyme Cathepsin-L is proteolytically processed in the lysosomal lumen to form a mature enzyme. To further examine the functionality of the lysosomes, we examined processing of Cathepsin-L and found no significant difference in Cathepsin-L processing in VCP-RNAi animals or animals treated with DBeQ (Figure 6—figure supplement 1B). Thus, disruption of the tubular lysosomal network does not appear to affect the proteolytic capacity of sarcoplasmic lysosomes.

To this point, our data suggest that loss of VCP disrupts the fusion of autophagosomes with functional lysosomes and, in parallel, causes the collapse of the tubular lysosomal network. Based on this, we expected to find evidence of failed autophagy in VCP knockdown muscle. In wild type muscles stained with a poly-ubiquitin antibody, we observed small puncta around the nucleus and a few small puncta in the cytoplasm (Figure 6A). These small puncta likely represent active sites of protein degradation by the proteasome. However, in muscles expressing VCP-RNAi we observed a dramatic accumulation of cytoplasmic poly-ubiquitin aggregates (Figure 6B). Even acute treatment with DBeQ was sufficient to produce cytoplasmic poly-ubiquitin aggregates (Figure 6C–E). We also note that VCP inhibition caused a dramatic decrease in poly-ubiquitin conjugates around the nucleus, which likely reflects failed delivery of poly-ubiquitinated proteins to the proteasome. Importantly, we find that Spin-positive lysosomes are devoid of poly-ubiquitin staining (Figure 6F), an observation that is consistent with our model of failed fusion of autophagosomes with lysosomes.

Figure 6 with 2 supplements see all
Disruption of the tubular auto-lysosomal network correlates with increased poly-Ubiquitin aggregates, impaired mitochondria and increased lipofuscin granules.

(A, B) Wild type (A) and VCP-RNAi (B) expressing muscles were fixed and stained with a poly-Ubiquitin antibody. Nuclei with localized poly-Ubiquitin staining are apparent in A. Nuclei are indicated (dashed circle) in B. (C, D) Wild type animals were treated with DMSO (C) or the VCP-specific inhibitor DBeQ (D), fixed and stained with a poly-Ubiquitin antibody. (E) Quantitation of the number of poly-Ubiquitin aggregates per 50 µm2 from wild type muscles treated with DMSO for 4 hr or DBeQ for various times (n = 9, *p < 0.05, **p < 0.01). (F) Localization of Spin-GFP and poly-Ubiquitin in muscles expressing VCP-RNAi. (G, H) Mitotracker-C2TMRos staining in control (G) and VCP-RNAi (H) muscles. (IK) Autofluoresence at 488 nm and lysotracker staining in wild type (I), muscles expressing VCP-RNAi (J), and wild type muscles treated with the VCP-specific inhibitor DBeQ for 4 hr (K).

https://doi.org/10.7554/eLife.07366.015

In addition to clearing protein aggregates from the cytoplasm, autophagy is also responsible for clearing damaged organelles, including mitochondria. We examined the functional pool of muscle mitochondria in vivo using mitotracker-Orange CM-H2TMRos, which selectively stains mitochondria with an active membrane potential. Muscles expressing VCP-RNAi displayed less mitotracker-Orange CM-H2TMRos staining and the visualized mitochondria had an altered morphology, appearing round and dispersed rather than being densely packed, tubular structures (Figure 6G,H). The swollen mitochondria observed in the VCP RNAi expressing muscles are likely defective and should be a prime target for mitophagy-dependent degradation. Taken together with the appearance of polyubiquitin aggregates, these data are consistent with an overall defect in autophagic clearance of proteins and defective organelles. However, since VCP has also been shown to be recruited directly to damaged mitochondria (Kim et al., 2013) we cannot rule out the possibility that the effects on mitochondrial morphology and membrane potential are due to direct VCP functions at the mitochondrial outer membrane.

Finally, we noted an accumulation of lipofuscin granules in VCP-RNAi expressing muscles. Lipofuscin granules are a conglomerate of polymerized non-degradable proteins and lipids that build up in the lysosomal lumen (Szweda et al., 2003). A distinguishing property of lipofuscin granules is that they exhibit auto-fluorescence at 488 nm. Wild type or VCP-RNAi muscles were stained with LysoTracker-Red and imaged at both 488 nm (green) and 555 nm (magenta). The wild type muscles did not display any detectable auto-fluorescence at 488 nm (Figure 6I). But, we observed strong auto-fluorescent puncta in VCP-RNAi that co-localized with LysoTracker (Figure 6J), indicative of lipofuscin granules. We did not observe autofluorescent puncta to the same extent when wild type muscles were treated with DBeQ for 4 hr (Figure 6K), suggesting that lipofuscin accumulation is a progressive phenotype. Alternatively, lipofuscin granule accumulation could require loss of VCP protein rather than just loss of VCP catalytic activity. These data suggest that maintaining the structural integrity of lysosome tubules is critical for lysosome function.

VCP-related disease mutations disrupt lysosome tubules

Finally, we asked whether overexpression of VCP transgenes that harbor disease-causing mutations impairs the presence or dynamics of tubular lysosomes. To date, a total of 19 missense mutations in 13 different residues are associated with IBMFD that reside in either the Cdc48 homology domain, the L1 linker domain or the D1 ATPase domain (Figure 7A and [Ju and Weihl, 2010]). We selected one mutation from each of these three domains to examine the effect on lysosome tubules: R155H, R191Q and A232E. R155 is the most common hereditary mutation and is located in the CDC48 homology domain, which is a protein interaction module that plays an important role in VCP substrate binding (Ju and Weihl, 2010). R191 is located in the linker region between the CDC48 domain and the first ATPase catalytic domain (D1). A232 is located at the beginning the of the D1 domain and is the most severe clinical mutation (Watts et al., 2004). All three residues are conserved in the Drosophila protein (Figure 7A). As a control, we show that over-expression of wild type dVCP does not alter tubular lysosomes (Figure 7B). We then demonstrate that over-expression of each of the mutant VCP transgenes profoundly impairs the tubular lysosomal network (Figure 7C–F). In parallel, we find an increase in sarcoplasmic auto-fluorescence when these VCP mutants are over-expressed (Figure 7B–G). Overexpression of the A229E mutation, which is the most severe clinical mutation, caused the largest increase in auto-fluorescence compared to the other clinical mutations (Figure 7F). Because over-expression of the disease relevant VCP transgenes phenocopies VCP-RNAi expression, these data suggest that the disease mutations are dominant interfering mutations. Thus, overexpression of VCP disease-associated mutations disrupts lysosome tubules in vivo, an effect that causes accumulation of cytoplasmic lipofuscin granules.

Pathogenic VCP alleles disrupt the tubular auto-lysosomal network.

(A) Schematic diagram of VCP protein. Top: Human pathogenic VCP mutations are labeled on the cartoon. Bottom: sequence alignment of human VCP and Drosophila VCP/Ter94 pathogenic mutant regions. (BF) Autofluoresence at 488 nm and lysotracker staining in wild type muscles expressing VCP-WT (B), VCP-RNAi (C) VCP-R152H (D), VCP-R188Q (E) or VCPA-229E (F) transgenes. (G) Quantitation of the number of auto-fluorescent puncta per 50 µm2 in the genotypes indicated (n = 9, *p < 0.05, **p < 0.01).

https://doi.org/10.7554/eLife.07366.018

Discussion

Here we demonstrate that lysosomes form a dynamic, tubular array that extends throughout the sarcoplasm of Drosophila muscle, in vivo. To our knowledge, this is the first observation of such an extensive, dynamic tubular lysosomal network in any in vivo system. We define this as a lysosomal network because it has a low pH, it is labeled by the late endosomal marker Spin-GFP, does not co-localize with the ER, golgi apparatus, early endosomes or recycling endosomes, and we find that mCherry-Atg8 traffics to this compartment, indicative of autophagosome fusion with a network of tubular lysosomes. Lysosomal network dynamics require an intact microtubule cytoskeleton, clathrin heavy chain, and VCP. We provide evidence that the catalytic activity of VCP is continuously required to sustain network integrity. When VCP is depleted or inhibited, the lysosomal network collapses and both cellular proteostasis and autophagy are compromised. Collapse of the network correlates with cellular manifestations of VCP-associated degenerative diseases, including the appearance of protein inclusion bodies (Kimonis et al., 2008; Weihl et al., 2009). While many questions remain unanswered, our discovery of tubular lysosome dysfunction following overexpression of pathogenic VCP mutants represents one the earliest markers of VCP-dependent muscle pathology. Further characterization could pave the way toward methods that might beneficially stabilize lysosomal function and ameliorate the progression of widespread degenerative disease.

Role of an extended tubular lysosome network in muscles

In most tissues, autophagy is induced upon nutrient starvation, but muscles are one of the unique tissues in which autophagy occurs in the absence of starvation (Mizushima et al., 2004). In fact, basal autophagy is required to maintain muscle mass (Masiero et al., 2009). The constitutive levels of autophagy in muscle are likely related to the large energy requirement of muscles compared to other cell types and their ability to serve as a source of metabolic energy for other organs (Sandri, 2010). Muscle sarcoplasms are also unique in that they are shared between multiple nuclei and are much larger in volume compared to many other cell types. We propose that the observed extended, lysosomal network that is maintained by VCP is a cellular solution to ensure highly efficient autophagy throughout the entirety of muscle sarcoplasms. It is as if the lysosomes vascularize the sarcoplasm, ensuring that no portion of the sarcoplasm is ever far from a lysosomal depot where autophagic cargo can be degraded. Likewise, this network could facilitate the local recycling of nutrients throughout the sarcoplasm to effectively meet local demands. A similar argument can be made regarding the autophagy-dependent turnover of mitochondria, which are densely distributed throughout the muscle. In support of this, VCP is critical for the rapid degradation of muscle proteins during muscle atrophy and expression of a dominant negative form of VCP reduces protein degradation by both proteasomes and lysosomes (Piccirillo and Goldberg, 2012).

Lysosomal membranes have been observed to extend tubules and fission off to produce de novo lysosomes following fusion of autophagosomes with lysosomes (Yu et al., 2010). This process, termed ALR, serves as a mechanism to recycle lysosomes when autophagic demand is high (Yu et al., 2010). It is unknown whether ALR is a constitutive process that participates in ongoing, steady-state proteostasis in vivo or whether it is a process that is specifically induced following stress-induced autophagy. It is possible that the extended, dynamic lysosomal tubule network that we observe is related to ALR, perhaps acting as a platform for efficient ALR throughout the sarcoplasm. In support of this, the mammalian Spinster homolog is required for ALR (Rong et al., 2011). However, the relatively low frequency with which we observed scission events and the formation of new small lysosome vesicles is not entirely consistent with this idea.

The tubular network identified in this study more closely resembles the lysosome tubular networks that have been observed in cultured macrophage cells (Swanson et al., 1987a, 1987b; Knapp and Swanson, 1990). Macrophage lysosome tubules can form a web of tubules that appear to be stably connected throughout the cytoplasm (Swanson et al., 1987b). The purpose of lysosome tubules in this system is still mysterious. We have now identified specific defects associated with loss of lysosome tubules and our data suggest that the tubular network may distribute auto-lysosomal activity throughout the cell. Clues to the function of tubular lysosomes might also be gleaned from studies of early endosomes. Early endosomes form vesicular-tubular structures similar to what we have observed here (Huotari and Helenius, 2011). In early endosomes it has been established that tubules can be discrete membrane compartments with a different lipid composition and cargo compared to that within the vesicular body of the early endosome (Huotari and Helenius, 2011). By analogy, the ability of lysosomes to exist in a tubular-vesicular state in cellular contexts where their functional demand is high, might allow lysosomes to execute diverse functions more efficiently through compartmentalization.

Consequences of tubular lysosome dysfunction

VCP and the yeast homologue Cdc48 have been ascribed many functions within the cell including cell cycle progression (Moir et al., 1982), UPS and ERAD protein degradation (Meyer et al., 2012; Wolf and Stolz, 2012), mitophagy (Taylor and Rutter, 2011) and classical autophagy (Ju et al., 2009; Tresse et al., 2010a; Dargemont and Ossareh-Nazari, 2012; Meyer et al., 2012). VCP achieves these diverse activities, in part, through its generalized function as an ubiquitin-dependent ‘segregase’ that dissociates protein conjugates tagged with ubiquitin from protein complexes and organelle membranes (Halawani and Latterich, 2006; Meyer et al., 2012). For each process that relies upon VCP activity, different cofactors control VCP localization and function (Baek et al., 2013). To this list of VCP-mediated activities, we now add the action of VCP in controlling the integrity and dynamics of a tubular lysosomal system and fusion of autophagosomes with tubular lysosomes. Additional, future experimentation will be necessary to determine which specific VCP-mediated molecular mechanism(s) is most directly relevant to the integrity and dynamics of lysosomal tubules in muscle.

The phenotypic consequences following the loss or inhibition of VCP in Drosophila muscle include the collapse of the tubular lysosomal network, failed fusion of autophagosomes with lysosomes, accumulation of sarcoplasmic poly-ubiquitin aggregates, accumulation of lipofuscin granules, impaired mitochondria and impaired muscle health. All of these phenotypes are hallmarks of degenerative diseases that are associated with mutations in VCP (Watts et al., 2004). We can now ascribe some of these phenotypes to the action of VCP at lysosomal membranes. Our data indicate that VCP localizes to autolysosomes and loss of VCP causes collapse of the tubular lysosomal network and failed autophagosome-lysosome fusion. Furthermore, when previously sequestered VCP is released back into the cytoplasm, VCP translocates to dormant autolysosomes and tubulation ensues. Although we cannot distinguish the role of VCP in the initiation and/or maintenance of lysosome tubules, these data argue that VCP acts at the autolysosmal membrane to control autolysosomal dynamics and the progression of autophagic protein clearance. If VCP is necessary for normal activity of the autophagy-lysosome system in muscle as our data suggests, then it seems likely that the accumulation of poly-ubiquitin aggregates and lipofuscin granules are a direct consequence of impaired VCP-dependent lysosomal function. This assertion is further supported by our demonstration that over-expression of pathogenic VCP mutant transgenes disrupt the tubular lysosomal network and also cause accumulation of ubiquitin and lipofuscin material. Importantly, these transgenes in Drosophila do not appear to disrupt VCP roles in the UPS or ERAD pathways, emphasizing the role of impaired autophagy in pathogenic VCP phenotypes (Tresse et al., 2010b; Chang et al., 2011). We acknowledge that lipofuscin granule accumulation following overexpression of pathogenic VCP mutant transgenes could represent a byproduct of proteotoxic stress caused by defects in the UPS or autophagy/lysosome pathways that eventually lead to the transformation of proteins into non-degradable products. Ultimately, more detailed biochemical analyses will be required to elucidate precisely how VCP functions at lysosome membranes and how this activity might be coordinated with other aspects of VCP function throughout the cell. Taken together, our identification of tubular lysosomes in Drosophila muscle challenges the traditional view of vesicular lysosomes and suggests that lysosome structures can be more versatile than previously assumed. Understanding how lysosomes regulate their morphological state will be an exciting avenue for future studies.

Materials and methods

Experimental procedures

Fly stocks

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The following transgenic fly stocks were generated in this study: UAS-Spin-RFP, UAS-Spin-GFP and UAS-Ter94/VCP-Venus. Drosophila spinster and ter94 cDNA were obtained by amplifying from DGRC (Drosophlia Genomics Resource Center, Bloomington, IN) clones AT25382 and GM02885 respectively. The cDNAs were cloned into the Gateway pENTR vector (Invitrogen, South San Francisco, CA) and subsequently cloned into destination vector pTWV obtained from the Drosophila Gateway Vector Collection (Carnegie Institution, Baltimore, MD). Transgenic animals were then generated by BestGene. UAS-mCherry-Atg8a and UAS-GFP-mCherry-Atg8a stocks were purchased from the Bloomington Stock Center. UAS-RNAi stocks were purchased from the Vienna Drosophila Resource Center. Pathogeneic UAS-Ter94 alleles were a kind gift from TK Sang (National Tsing Hua University, Hsinchu, Taiwan). Golgi markers UAS-ManII-GFP and UAS-GalT-YFP were a kind gift from B Ye (University of Michigan, Ann Arbor, MI). Gal4 muscle drivers used in this study include: BG57-Gal4 and MHC-Gal4.

Microscopy methods

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Third instar larvae were dissected in Schneider's insect cell media (Gibco, Grand Island, NY) supplemented with 10% FBS (Gibco) and Penicillin/Streptomycin (Gibco) and all live imaging was performed in insect cell media. For all imaging experiment, at least 3 muscles in 3 animals were imaged to account for variances between muscles and animals and the most representative images are shown in each figure (n ≥ 9). The following drugs were diluted in insect cell media to the following final concentrations: 1uM LysoTracker Red DND-99 (Life Technologies, South San Francisco, CA), 1uM ER Tracker green (Life Technologies), 10uM Nocodazole (Sigma, St. Louis, MO), 10uM Latrunculin A (Life Technologies), 10uM DBeQ (Sigma), 1uM MG132 (Sigma), 100 nM Concanamycin A (Santa Cruz Biotechnology, Dallas, TX), 2ug/ml TM (Sigma) and 500 nM Mitotracker-Orange-CM-H2TMRos (Life Technologies). LysoTracker, ER Tracker and Mitotracker-Orange-CM-H2TMRos were incubated on the dissected larvae for 1 hr prior to imaging. Nocodazole, LatA, DBeQ, TM, Concanamycin A and MG132 were incubated on the dissected larvae for 3–4 hr prior to imaging.

For Ubiquitin staining, larvae were dissected and fixed with 4% PFA for 15 min. After fixation, larvae were washed 4× with PBS-T and incubated with anti-poly-Ubiquitin (Thermo Scientific, Waltham, MA) at 1:1000 dilution overnight at 4°C. Larvae were washed again 4× with PBS-T, incubated with a fluorescently labeled secondary antibody at 1:5000 dilution (Life Technologies), and washed again 4× with PBS-T before mounting on a slide with vectashield for imaging.

Imaging was performed on an inverted Axiovert 200 microscope (Zeiss) using a 100× Plan Apochromat objective (1.4NA). Images were captured with a CoolSnap HQ2 CCD camera (Photometrics) and de-convolved using Slidebook 5.0 software (Intelligent imaging innovations, Denver, CO). Image quantification was performed with imageJ software (NIH). Volume rendering was performed with Slidebook 5.0 software. Any adjustment of brightness or contrast was performed using Slidebook 5.0 software, and always applied to the entire image.

Western blotting methods

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Third instar larvae were dissected and muscle preparations were immediately transferred into 5× SDS sample buffer and denatured by boiling for 10 min. Proteins were resolved by SDS-PAGE on a 4–12% Bis-Tris gel (Life Technologies), transferred to a nitrocellulose membrane, immunoblotted with primary and HRP-conjugated secondary antibodies (Life Technologies) and detected using an ECL chemi-luminescent reagent (Life Technologies). The following primary antibodies were used: anti-VCP (Cell Signaling, Danvers, MA) at 1:1000, anti- GRP78/HSPA5 (Thermo Scientific) at 1:2500, insect anti-Cathepsin L (R&D Systems, Minneapolis, MN) at 1:1000, and anti-Tubulin E7-c (Developmental Studies Hybridoma Bank, University of Iowa, IA) at 1:10,000. Secondary HRP conjugated antibodies (GE Lifesciences, Pittsburgh, PA) were used at 1:5000.

Electrophysiology methods

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Sharp-electrode recordings were made from muscle 6 in abdominal segments 2 and 3 from third-instar larvae using an Axoclamp 900A amplifier (Molecular Devices), as described previously (Frank et al., 2006). Recordings were made in HL3 saline containing the following components: NaCl (70 mM), KCl (5 mM), MgCl2 (10 mM), NaHCO3 (10 mM), sucrose (115 mM), trehalose (5 mM), HEPES (5 mM), and CaCl2 (0.3 mM). Mean EPSP, mEPSP amplitude, and Rin were obtained by averaging values across all NMJs for a given genotype. EPSP traces were analyzed with custom-written routines in MATLAB (Mathworks, Natick, MA) as previously described (Gaviño et al., 2015). mEPSP traces were analyzed in IGOR Pro 6.3 (Wave-Metrics; custom script submitted with this manuscript).

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Decision letter

  1. K VijayRaghavan
    Reviewing Editor; National Centre for Biological Sciences, Tata Institute for Fundamental Research, India

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for sending your work entitled “VCP-Dependent Muscle Degeneration is Linked to Defects in a Dynamic Tubular Lysosomal Network in vivo” for consideration at eLife. Your article has been favorably evaluated by K VijayRaghavan (Senior editor and Reviewing editor) and three reviewers.

In the current manuscript, Johnston et al. identify a tubular network in the muscle of Drosophila, which appears to arise from the fusion of multiple lysosomal vesicles. This remarkable architecture of the lysosomal system requires the AAA-ATPase Valosyl Containing Protein (VCP), mutations of which are implicated in human degenerative disease. Deletion of VCP (or its replacement with disease-relevant mutants) results in disassembly of the lysosomal tubular network and in a series of defects, including defective autophagy and poly-ubiquitin aggregation, linking this unique architecture of the lysosomal system to maintenance of cellular quality control.

Overview of referee comments:

Based on the overall evaluation and comments arising from the collective reviews, we first outline the broad areas of concern, which can be binned into two key points.

A) The referees agree that the novelty that makes this manuscript potentially suitable for eLife is the tubular organisation of the lysosomal network in muscle. A more thorough characterisation of this striking sub-cellular structure will therefore be valuable and necessary. Better quantification and explanation of sample sizes as well as controls are needed to demonstrate that the observation of the lysosomal phenotypes seen are not biased by choice of probes used (Spinster vs. Lysotracker for example). These will much strengthen the characterisation of the cellular network and the authors could consider using additional markers and co-localisation studies.

B) Each reviewer had concerns about the uncertainty underlying mechanisms whereby VCP exerts effects on lysosome morphology and/or function. Does VCP separately regulate lysosome tubulation, the fusion of autophagosomes with lysosomes and/or some other critical function of lysosomes?

We do see that addressing the first set of points (A) appears very feasible in a timely manner. Answers to these queries would likely solidify but probably not grossly change the current set of morphological observations. Dissecting out the issue of mechanism is likely to be beyond what the authors could hope to achieve in a timely manner. While we feel that 'descriptive' findings of the paper are actually of sufficient interest to be published in eLife in the absence of further experiments to demonstrate 'mechanisms', we are concerned that the final version of the paper should be careful not to overreach in claiming more mechanistic insight than is supported by the actual data. If the data cannot distinguish between multiple putative lysosomal actions for VCP, then the paper should state and reflect this. The authors have been reasonably careful in this regard, but this is just a flag that the revisions should continue to adhere to this path.

Simply discussing the concerns expressed in (B) above should be sufficient and we do not require experiments in this direction.

Main comments:

The demonstration that lysosomes in Drosophila muscles exhibit a very dramatic tubular morphology is unexpected, intriguing and new. The suggestion is that the collapse of this tubular network of lysosomes might represent a mechanism through which impaired valosin containing protein (VCP) function contributes to human disease. This collapse of the lysosomal tubular network is accompanied by defects in autophagosome-lysosome fusion and the accumulation of lipofuscin within lysosomes. VCP is best characterized as an ATPase that facilitates the degradation of ubiquitinated proteins and is vaguely linked to diverse other cellular processes. The authors build on their new observations to suggest a novel role for VCP in promoting either lysosome tubulation, the fusion of autophagosomes with lysosomes and/or some other critical function of lysosomes. While each of these possibilities is reasonable based on the available data, the manuscript ultimately suffers from the lack of a clear elucidation of the specific contribution of VCP to maintenance of lysosome function. Does VCP contribute to one specific aspect of lysosome function that when disrupted leads to the various phenotypes characterized in this study? Or do these phenotypes reflect multiple distinct functions of VCP? While the tubular morphology of lysosomes in muscle represents a striking new observation and the dissemination of this knowledge has great value, however, it is not clear that the disruption of lysosomal tubulation by VCP depletion/mutations underlies the defects in lysosome function (autophagosome fusion and lipofuscin accumulation) that are simultaneously observed under such conditions. Thus, the major limitation is the lack of a precise elucidation of the function of VCP at lysosomes. The authors should keep in mind the comments in the overview, and here, to prioritise how to address this 'main comment'.

Specific points to be addressed:

1) While the images of this lysosomal network are striking and the description of the dynamics is succinct, there is hardly any quantitative statement in their analyses, or in experimental procedures e.g. unclear how many larvae prep were imaged in each experiment. Figure 2E-G: how penetrant of the rescue of VCP(RNAi) by hVCP is? Generally, based on what criteria, a transgene is over-expressed?

2) In the first paragraph of the subsection “VCP is required for the integrity and dynamics of the extended tubular lysosomal network”: The authors state that they pursued a candidate-based RNAi screen, but the list of candidates is missing. Do they want to list them here, or leave it out?

3) It is still somewhat confusing regarding VCP's role in this lysosomal network formation or maintenance. The Phalloidin staining of muscle of VCP-RNAi in Figure 6–figure supplement 1 also seems to show quite noticeable muscle structural defects.

4) Regarding the analyses of the disease-associated mutant VCP, are these mutations loss of function or gain of function? Can they analyse these mutant VCP using the VCP(RNAi) plus hVCP transgene expression scheme as in Figure 2?

5) In the second paragraph of the subsection “Drosophila sarcoplasmic lysosomes form an extended dynamic tubular array”, while the lysotracker staining supports the Spin-RFP structure is not an artifact, it remains unclear whether over-expressing Spin causes expansion or distortion of this network as the images in Figure 1D and E do look different. It would seem that they can resolve this issue by lysotracker staining in Spin-RFP or Spin-GFP lines.

6) In the third paragraph of the subsection “Autophagosome membranes co-localize with the tubular lysosome network” and Figure 3D-E, the GFP-mcherry-Atg8 reporter is very clever. This construct should be referenced and shown the figure. The authors may want to mention that Spin-GFP is not quenched due to topology of the protein.

7) VCP has been previously implicated in the maturation of autophagosomes (Ju et al., JCB, 2009; Tresse et al., Autophagy, 2010). Also in the endosomal sorting of cargos for lysosomal degradation (Ritz et al., NCB, 2011). This new manuscript essentially confirms these results in the Drosophila muscle but do not offer further significant insight into mechanism.

8) The specific defect in lysosomes that arises in VCP depleted cells remains vague. Normal cathepsin L maturation and lysotracker labeling suggest that some basic functions are retained. Nonetheless, the accumulation of lipofuscin indicates a defect. More insight into the specific lysosome defect would be helpful in nailing down the actual function of VCP at lysosomes (this might be beyond the scope of the current manuscript, see Overview above).

9) It is not clear why the alterations in mitochondrial morphology in VCP depleted cells might reflect an autophagy defect.

10) The authors identify the tubular network as lysosomal based on the localization of Spinster, a transporter previously identified on lysosomal vesicles, as well as staining of the tubular network with lysotracker (Figure 1). However, to fully confirm that lysosomes are the main (and only) constituents, other markers should be examined, including LAMP1/2 and Cathepsins. They should also show that bona fide Golgi and ER markers do not co-localize with Spin-RFP, and clarify the spatial relationship of these compartments to the tubular network.

11) What is the ultrastructural appearance of this network? Is it electron-dense, as are classical lysosomes? Does it contain multi-vesicular body-like objects? Is its limiting membrane single or double? Although EM-grade fixation may disrupt network architecture, an EM analysis of the resulting vesicles could still yield useful information about their composition and origin. The authors may want to consider whether this experimental avenue can be speedily explored.

12) Staining of the tubular network with lysotracker, a dye that accumulates in acidic compartments, supports that this network is primarily composed of acidic organelles (Figure 1E). The vacuolar H+ ATPase (V-ATPase) mediates acidification of lysosomes. Do V-ATPase inhibitors such as concanamycin and Bafilomycin A dissipate the internal acidity of the network? We feel that these experiments should suffice to demonstrate the importance of acidification. However, regarding pH, while lysotracker suggests that this compartment is acidic, it does not allow a precise determination of pH values. Precise measurement of pH may be technically challenging and documenting regional differences not of significant additional value. But, if feasible and the authors wish to embark on this direction, they could employ ratiometric GFP constructs or, alternatively, Lysosensor dyes. By building a calibration curve and interpolating the value, they should be able to come up with a precise value (possibly matching the lysosomal internal pH, which is within the 4.0-5.0 range) and also determine whether pH is homogeneous throughout the network or if subregions with different acidity values exist.

13) The autophagic marker Atg8 is distributed along the length of the tubular network, albeit only in certain subregions (Figure 3A). The authors propose that these are regions of ongoing autophagosome-lysosome fusion, leading to progressive Atg8 degradation. One important question is whether this continuous influx of autophagic membranes may in fact enable the accretion of the lysosomal tubular network. The authors should knock down key autophagic mediators such as Atg5 or Atg7, and measure the impact on the morphology of Spns-RFP positive vesicles.

14) Treatment with the VCP inhibitor DbEQ and with the proteasome inhibitor MG132 leads to the formation of VCP-GFP positive rod-like structures and an overall increase in cellular VCP levels (Figure 4D, E). Thus, under normal circumstances VCP may be subjected to rapid degradation. The authors should verify this point by quantifying total VCP-GFP fluorescence as well as by western blotting.

15) Treatment with MG132 causes the dissolution of the tubular network (Figure 4E). This effect is attributed to the sequestration of VCP in rod-like structures, which would block its action in a dominant-negative fashion. However, the link here is tenuous given that many proteins undergoing proteasome-regulated turnover could be involved in the tubular network architecture. Another way to test this point is to induce VCP aggregation in a proteasome-independent way using protein homo- or hetero-dimerization systems (i.e. FKBP-FRB) to see whether a similar effect is obtained. If successful, the authors should use this technique to verify that the same phenotypes observed upon RNAi-mediated suppression of VCP (poly-Ub aggregates, mitochondrial damage) also occur.

16) An important question concerns the mechanism of action of VCP in lysosomal network maintenance, autophagosome-lysosome fusion etc. Are these effects due to a direct action of VCP on lysosomal and autophagosomal proteins? Is it an indirect effect of VCP loss of function due, for instance, to ER stress? Does induction of ER stress by other means result in the same effects?

https://doi.org/10.7554/eLife.07366.019

Author response

Based on the overall evaluation and comments arising from the collective reviews, we first outline the broad areas of concern, which can be binned into two key points.

A) The referees agree that the novelty that makes this manuscript potentially suitable for eLife is the tubular organisation of the lysosomal network in muscle. A more thorough characterisation of this striking sub-cellular structure will therefore be valuable and necessary. Better quantification and explanation of sample sizes as well as controls are needed to demonstrate that the observation of the lysosomal phenotypes seen are not biased by choice of probes used (Spinster vs. Lysotracker for example). These will much strengthen the characterisation of the cellular network and the authors could consider using additional markers and co-localisation studies.

We appreciate the reviewers’ comments, which we have addressed with new experiments and text revisions. These changes are briefly listed here and are further detailed in the responses to specific criticisms (below). Collectively, these additional experiments provide a more thorough characterization of the tubules and more clearly define these tubules as lysosomal. Data for items A1-A3 are now reported in new Figure 2.

A1) We performed experiments to co-label the lysosomal tubules with Spinster-GFP and Lysotracker in the same muscle. We show near perfect co-localization between Spinster-GFP and Lysotracker, demonstrating that these probes are marking the same tubular organelle structures.

A2) We have co-imaged Spin-RFP labeled lysosomal tubules with several additional markers including ER tracker, medial and trans golgi markers as well as markers for both early and recycling endosomes. We also performed co-imaging of Spin-RFP with a mitochondrial marker. All of these markers show distinct localization patterns compared to Spin-RFP labeled tubules. As a note to the reviewers, golgi organization in skeletal muscle is distinct from the classical golgi stacks observed in many cells. Vertebrate muscle shows a unique, distributed, vesicular organization of the golgi throughout the muscle (Ralston et al., J. Neurosci., 2001). We find that this organization is conserved in Drosophila muscle.

A3) We performed an additional experiment to test the acidification of Spin-labeled tubules. We treated muscles with the V-ATPase specific inhibitor concanamycin A and observed a decrease in acidity in the tubules. ConA inhibits the lysosomal V-ATPase, verifying that the Spin-positive tubules are acidic and lysosomal.

A4) We have modified the text to clarify sample sizes and our approach to the presentation of data. Regarding the issue of sample size. Tubules are present in every muscle that we have examined. Thus, phenotypic penetrance is essentially 100%. In addition, we went to the extent of imaging these tubules in the live, un-dissected animal (an experiment reported in the original submission). This was done to verify that the presence of tubules is not an artifact of animal dissection. We see tubules in all muscle cells in the live organism. We have added a statement to the text to this effect (Results section): “Tubules were observed in every muscle and there were no apparent differences in tubule abundance or architecture between different muscles.” We have included samples sizes in figure legends where necessary to interpret statistical comparisons. We also now state in the Methods section that at least 3 muscles in 3 animals (N≥9) were imaged and the most representative images are shown (subsection “Microscopy methods”). Regarding the collection of live imaging data for movies, more than 10 muscles were imaged (N≥10) for each experiment. In most instances, many more muscles than this were imaged for our experiments.

B) Each reviewer had concerns about the uncertainty underlying mechanisms whereby VCP exerts effects on lysosome morphology and/or function. Does VCP separately regulate lysosome tubulation, the fusion of autophagosomes with lysosomes and/or some other critical function of lysosomes?

We do see that addressing the first set of points (A) appears very feasible in a timely manner. Answers to these queries would likely solidify but probably not grossly change the current set of morphological observations. Dissecting out the issue of mechanism is likely to be beyond what the authors could hope to achieve in a timely manner. While we feel that 'descriptive' findings of the paper are actually of sufficient interest to be published in eLife in the absence of further experiments to demonstrate 'mechanisms', we are concerned that the final version of the paper should be careful not to overreach in claiming more mechanistic insight than is supported by the actual data. If the data cannot distinguish between multiple putative lysosomal actions for VCP, then the paper should state and reflect this. The authors have been reasonably careful in this regard, but this is just a flag that the revisions should continue to adhere to this path.

Simply discussing the concerns expressed in (B) above should be sufficient and we do not require experiments in this direction.

We thank the reviewers for their consideration and for their enthusiasm regarding our basic findings. Indeed, we hope that our work will spur new ideas and experimentation on VCP activity in muscle, which is one of the primary tissues compromised in human patients harboring disease-causing mutations in VCP. We have made some changes to the text, particularly in the Discussion, to make sure that we are appropriately circumspect regarding the mechanism of action of VCP.

Main comments:

The demonstration that lysosomes in Drosophila muscles exhibit a very dramatic tubular morphology, is unexpected, intriguing and new. The suggestion is that the collapse of this tubular network of lysosomes might represent a mechanism through which impaired valosin containing protein (VCP) function contributes to human disease. This collapse of the lysosomal tubular network is accompanied by defects in autophagosome-lysosome fusion and the accumulation of lipofuscin within lysosomes. VCP is best characterized as an ATPase that facilitates the degradation of ubiquitinated proteins and is vaguely linked to diverse other cellular processes. The authors build on their new observations to suggest a novel role for VCP in promoting either lysosome tubulation, the fusion of autophagosomes with lysosomes and/or some other critical function of lysosomes. While each of these possibilities is reasonable based on the available data, the manuscript ultimately suffers from the lack of a clear elucidation of the specific contribution of VCP to maintenance of lysosome function. Does VCP contribute to one specific aspect of lysosome function that when disrupted leads to the various phenotypes characterized in this study? Or do these phenotypes reflect multiple distinct functions of VCP? While the tubular morphology of lysosomes in muscle represents a striking new observation and the dissemination of this knowledge has great value, however, it is not clear that the disruption of lysosomal tubulation by VCP depletion/mutations underlies the defects in lysosome function (autophagosome fusion and lipofuscin accumulation) that are simultaneously observed under such conditions. Thus, the major limitation is the lack of a precise elucidation of the function of VCP at lysosomes. The authors should keep in mind the comments in the overview, and here, to prioritise how to address this 'main comment'.

We agree that we have not pinpointed the specific activity of VCP that controls the integrity of the lysosomal tubule network in muscle. We thank the reviewers for acknowledging that doing so will be a major effort for the future, given the diverse functions that have been ascribed to VCP in the cell. Although the experiments required to elucidate the specific contributions of VCP on lysosome tubulation are beyond the scope of the present study, our findings expose a new avenue of VCP biology that is likely to spur many more studies in this direction. None-the-less, in response to this criticism, we have strengthened our treatment of VCP in our Discussion section. We now state, “VCP and the yeast homologue Cdc48 have been ascribed many functions within the cell including cell cycle progression (Moir et al., 1982), UPS and ERAD protein degradation (Meyer et al., 2012; Wolf and Stolz, 2012), mitophagy (Taylor and Rutter, 2011) and classical autophagy (Dargemont and Ossareh-Nazari, 2012; Ju et al., 2009; Meyer et al., 2012; Tresse et al., 2010a)… To this list of VCP-mediated activities we now add the action of VCP in controlling the integrity and dynamics of a tubular lysosomal system and fusion of autophagosomes with tubular lysosomes. Additional, future experimentation will be necessary to determine which specific VCP-mediated molecular mechanism(s) is most directly relevant to the integrity and dynamics of lysosomal tubules in muscle” (subsection “Consequences of tubular lysosome dysfunction”).

Specific points to be addressed:

1) While the images of this lysosomal network are striking and the description of the dynamics is succinct, there is hardly any quantitative statement in their analyses, or in experimental procedures e.g. unclear how many larvae prep were imaged in each experiment. Figure 2E-G: how penetrant of the rescue of VCP(RNAi) by hVCP is? Generally, based on what criteria, a transgene is over-expressed?

We have modified the text to clarify sample sizes and our approach to the presentation of data. Regarding the issue of sample size. Tubules are present in every muscle that we have examined. Thus, phenotypic penetrance is essentially 100%. In addition, we went to the extent of imaging these tubules in the live, un-dissected animal (an experiment reported in the original submission). This was done to verify that the presence of tubules is not an artifact of animal dissection (please see point A4 above).

For rescue experiments, the phenotypic characterization is all or none; the tubules are absent in the knockdown and present in the rescue muscle. The phenotype was scored in N≥10 rescue animals. We did not attempt to make any quantitative estimate of the magnitude of the rescue as the phenotype was an all or none assessment with ∼100% penetrance. We modified the results to say “Over-expressing human VCP in dVCPRNAi muscles rescued the formation of lysosome tubules in every muscle” to clarify (subsection “VCP is required for the integrity and dynamics of the extended tubular lysosomal network”).

2) In the first paragraph of the subsection “VCP is required for the integrity and dynamics of the extended tubular lysosomal network”: The authors state that they pursued a candidate-based RNAi screen, but the list of candidates is missing. Do they want to list them here, or leave it out?

This screen is still ongoing. As such, we would prefer to leave out the identification of the other hits that we are finding. We thank the editors for their understanding.

3) It is still somewhat confusing regarding VCP's role in this lysosomal network formation or maintenance. The Phalloidin staining of muscle of VCP-RNAi in Figure 6–figure supplement 1 also seems to show quite noticeable muscle structural defects.

Based on our results, we cannot distinguish between VCP’s role in formation or maintenance and we have added a sentence in the Discussion to make this point more clear (“Although we cannot distinguish the role of VCP in the initiation and/or maintenance of lysosome tubules, these data argue that VCP acts at the autolysosmal membrane to control autolysosomal dynamics and the progression of autophagic protein clearance”). We agree that the muscles are visibly degrading in VCP RNAi animals. But, other degenerative mutants (Figure 3) do not cause disruption of the lysosome network, indicating that the loss of the tubular network is not merely a byproduct of muscle degeneration and is a specific defect of VCP loss. This is stated in the Results section (subsection “VCP is required for the integrity and dynamics of the extended tubular lysosomal network”).

4) Regarding the analyses of the disease-associated mutant VCP, are these mutations loss of function or gain of function? Can they analyse these mutant VCP using the VCP(RNAi) plus hVCP transgene expression scheme as in Figure 2?

VCP forms a hexamer and over-expressing the disease mutations in the context of wildtype VCP produces a dominant-negative effect. It is not clear whether over-expression of human-VCP would provide much additional information and we have not pursued this further. However, given that VCP RNAi produces a phenotype that is similar to the over-expression of mutant VCP, our results imply that the VCP mutants have a loss-of-function effect on lysosomal tubule integrity. This point has been added to the end of the Results section.

5) In the second paragraph of the subsection “Drosophila sarcoplasmic lysosomes form an extended dynamic tubular array”, while the lysotracker staining supports the Spin-RFP structure is not an artifact, it remains unclear whether over-expressing Spin causes expansion or distortion of this network as the images in Figure 1D and E do look different. It would seem that they can resolve this issue by lysotracker staining in Spin-RFP or Spin-GFP lines.

It is our impression that the apparent difference between Lysotracker staining and Spin-RFP (or Spin-GFP) is caused by the intensity of the different labels; a membrane-localized RFP versus membrane permeable dye. It remains possible that there is a distorting effect of Spin-RFP or –GFP on the tubule network, but this seems tangential to the main line of investigation for this paper. We have now shown that Lysotracker co-localizes with Spin-GFP, so we are certain that these reagents label the same compartment. These data are now included in new Figure 2.

6) In the third paragraph of the subsection “Autophagosome membranes co-localize with the tubular lysosome network” and Figure 3D-E, the GFP-mcherry-Atg8 reporter is very clever. This construct should be referenced and shown the figure. The authors may want to mention that Spin-GFP is not quenched due to topology of the protein.

We have added the reference for the GFP-mCherry-Atg8a reporter in the text (subsection “Autophagosome membranes co-localize with the tubular lysosome network”). We have also added a sentence and reference to clarify that Spin-GFP is not quenched because the C-terminal GFP tag resides on the cytoplasmic side of the lysosome membrane.

7) VCP has been previously implicated in the maturation of autophagosomes (Ju et al., JCB, 2009; Tresse et al., Autophagy, 2010). Also in the endosomal sorting of cargos for lysosomal degradation (Ritz et al., NCB, 2011). This new manuscript essentially confirms these results in the Drosophila muscle but do not offer further significant insight into mechanism.

As stated above, we have added new text to our Discussion acknowledging that it will take considerable future experimentation to pinpoint the specific activities of VCP that control the integrity of the lysosomal tubule network in muscle. I believe that we have also been careful to acknowledge and cite relevant previous studies including Ju et al., (2009) and Tresse et al., (2010). We are grateful to the editors for acknowledging that we also make an experimental advance through the identification and characterization of a tubular lysosomal network in muscle that requires VCP and is influenced by disease-causing mutations in VCP.

8) The specific defect in lysosomes that arises in VCP depleted cells remains vague. Normal cathepsin L maturation and lysotracker labeling suggest that some basic functions are retained. Nonetheless, the accumulation of lipofuscin indicates a defect. More insight into the specific lysosome defect would be helpful in nailing down the actual function of VCP at lysosomes (this might be beyond the scope of the current manuscript, see Overview above).

We believe that it is beyond the scope of this study and thank the editors for acknowledging that this might be the case.

9) It is not clear why the alterations in mitochondrial morphology in VCP depleted cells might reflect an autophagy defect.

Damaged mitochondria commonly exhibit altered morphology and membrane potential. Because damaged mitochondria are typically cleared though mitophagy, the accumulation of mitochondria with altered morphology suggests a defect in mitochondrial clearance. We have included additional text in the corresponding section of the results to clarify this point (“The swollen mitochondria observed in the VCP RNAi expressing muscles are likely defective and should be a prime target for mitophagy-dependent degradation”).

10) The authors identify the tubular network as lysosomal based on the localization of Spinster, a transporter previously identified on lysosomal vesicles, as well as staining of the tubular network with lysotracker (Figure 1). However, to fully confirm that lysosomes are the main (and only) constituents, other markers should be examined, including LAMP1/2 and Cathepsins. They should also show that bona fide Golgi and ER markers do not co-localize with Spin-RFP, and clarify the spatial relationship of these compartments to the tubular network.

We have done our best to address the reviewers’ comments with available reagents and have added an entire new figure of data to our manuscript (new Figure 2). First, we have performed several of the suggested experiments. We co-imaged Spin-RFP with several other previously characterized organelle markers. Co-imaging Spin-RFP with previously characterized markers of mitochondria (mito tracker), ER (ER tracker), medial golgi (ManII), trans golgi (galT), early endosomes (Rab5) and recycling endosomes (Rab11) clearly demonstrates that Spin-RFP labeling is distinct from all of these organelles. These data are now presented in a new figure (Figure 2). We thank the reviewers for prompting us to pursue these experiments. These data significantly extend our understanding of the tubular lysosomal network within the complex muscle sarcoplasm. As a note to the reviewers, golgi organization in skeletal muscle is distinct from the classical golgi stacks observed in many cells. Vertebrate muscle shows a unique, distributed, vesicular organization of the golgi throughout the muscle (Ralston et al., J. Neurosci., 2001). Our data show that this organization is conserved in Drosophila muscle.

Unfortunately, we were unable to perform some of the other experiments suggested by the reviewers. First, to our knowledge, fluorescent probes for Cathepsins in Drosophila do not exist and the process of generating new reagents and verifying their activity would be beyond the stated scope of eLife reviews. We also attempted to co-localize Spin-RFP with Lamp1-GFP, but found that the currently available Lamp1-GFP constructs do not express well in muscles using either of two different muscle specific Gal4 drivers (MHC-Gal4 or BG57-Gal4). However, the reviewers should be aware that when Spin is expressed in HeLa cells, Spin-GFP co-localizes with Lamp1 (Sweeney et al., 2001). Furthermore, we show strong co-localization between muscle expressed Spin-RFP and Lysotracker (Figure 2A). We also provide new evidence that the lysosomal vATPase is necessary to maintain tubule acidification, consistent with lysosomal identify. When taken into consideration with additional information described below, there is considerable evidence that the tubular network in muscle is lysosomal.

11) What is the ultrastructural appearance of this network? Is it electron-dense, as are classical lysosomes? Does it contain multi-vesicular body-like objects? Is its limiting membrane single or double? Although EM-grade fixation may disrupt network architecture, an EM analysis of the resulting vesicles could still yield useful information about their composition and origin. The authors may want to consider whether this experimental avenue can be speedily explored.

These are all excellent questions (electron density, limiting membrane, etc.). We have performed extensive TEM on Drosophila muscle. But, without an independent electron-dense marker (immuno-gold or HRP) there is no way for us to be sure about the identification of these lysosomal structures. Working out the details by which we can unambiguously identify lysosomes in muscle is beyond the scope of the current work, but something that we wish to pursue in future studies.

12) Staining of the tubular network with lysotracker, a dye that accumulates in acidic compartments, supports that this network is primarily composed of acidic organelles (Figure 1E). The vacuolar H+ ATPase (V-ATPase) mediates acidification of lysosomes. Do V-ATPase inhibitors such as concanamycin and Bafilomycin A dissipate the internal acidity of the network? We feel that these experiments should suffice to demonstrate the importance of acidification. However, regarding pH, while lysotracker suggests that this compartment is acidic, it does not allow a precise determination of pH values. Precise measurement of pH may be technically challenging and documenting regional differences not of significant additional value. But, if feasible and the authors wish to embark on this direction, they could employ ratiometric GFP constructs or, alternatively, Lysosensor dyes. By building a calibration curve and interpolating the value, they should be able to come up with a precise value (possibly matching the lysosomal internal pH, which is within the 4.0-5.0 range) and also determine whether pH is homogeneous throughout the network or if subregions with different acidity values exist.

We have examined the effect of ConA treatment on GFP-mCherry-Atg8a fluorescence and localization in muscles. Under basal conditions, GFP-mCherry-Atg8a localizes to tubules, but the GFP fluorescence is quenched inside the tubules due to the acidic environment and only mCherry fluorescence is observed. However, upon treatment of ConA for 3 hours, we observed GFP and mCherry tubule labeling. These data reveal several important aspects of the lysosome tubules. First, it verifies by another method that the tubules are indeed acidic. Second, it indicates that the acidity of the tubules is dependent upon the V-ATPase, a lysosomal-specific proton pump. Finally, these data show that acidification of the lysosomes is not required to maintain the tubular architecture. These new data have been added to Figure 5.

We agree that quantitative exploration of the pH within the lysosomal network and the possibility that regional differences exist would be very interesting. However, we believe that it is beyond the scope of the current work.

13) The autophagic marker Atg8 is distributed along the length of the tubular network, albeit only in certain subregions (Figure 3A). The authors propose that these are regions of ongoing autophagosome-lysosome fusion, leading to progressive Atg8 degradation. One important question is whether this continuous influx of autophagic membranes may in fact enable the accretion of the lysosomal tubular network. The authors should knock down key autophagic mediators such as Atg5 or Atg7, and measure the impact on the morphology of Spns-RFP positive vesicles.

We have done the suggested experiment and find that inhibition of Atg7 by RNAi (a previously characterized RNAi line) does not disrupt the lysosome tubular network. Thus, continuous autophagic flux is not necessary for the integrity of the lysosomal network. These data have been added to Figure 4.

14) Treatment with the VCP inhibitor DbEQ and with the proteasome inhibitor MG132 leads to the formation of VCP-GFP positive rod-like structures and an overall increase in cellular VCP levels (Figure 4D, E). Thus, under normal circumstances VCP may be subjected to rapid degradation. The authors should verify this point by quantifying total VCP-GFP fluorescence as well as by western blotting.

We would like to clarify that we do not think cellular VCP levels increase upon treatment with MG132 or DBeQ. This is based on our observation that upon washout of MG132, the VCP rods dissolve and the cytoplasmic pool of VCP is restored to resting state. None-the-less, we followed the reviewers’ suggestion and examined total VCP levels upon treatment with MG132 or DBeQ by western blot analysis and did not observe any increase in total VCP levels. We have added these data to Figure 5.

15) Treatment with MG132 causes the dissolution of the tubular network (Figure 4E). This effect is attributed to the sequestration of VCP in rod-like structures, which would block its action in a dominant-negative fashion. However, the link here is tenuous given that many proteins undergoing proteasome-regulated turnover could be involved in the tubular network architecture. Another way to test this point is to induce VCP aggregation in a proteasome-independent way using protein homo- or hetero-dimerization systems (i.e. FKBP-FRB) to see whether a similar effect is obtained. If successful, the authors should use this technique to verify that the same phenotypes observed upon RNAi-mediated suppression of VCP (poly-Ub aggregates, mitochondrial damage) also occur.

Certainly, proteasome inhibition has many effects on the cell. However, we want to emphasize that the point of this experiment was not to implicate proteasome activity in the maintenance of tubular network architecture. Rather, the discovery that MG132 caused VCP aggregation was a fortuitous observation that allowed us to do a recovery experiment. We used MG132 to induce sequestration of VCP into cytoplasmic aggregates. Then, we were able to wash out MG132 and watch the effect(s) of re-establishing a cytoplasmic pool of soluble VCP. It is clear from the imaging data that VCP is sequestered from the cytoplasm during VCP aggregation and that cytoplasmic VCP is restored upon MG132 washout and tubulation was restored.

16) An important question concerns the mechanism of action of VCP in lysosomal network maintenance, autophagosome-lysosome fusion etc. Are these effects due to a direct action of VCP on lysosomal and autophagosomal proteins? Is it an indirect effect of VCP loss of function due, for instance, to ER stress? Does induction of ER stress by other means result in the same effects?

We have done the suggested experiment to test for the effects of ER stress. We induced ER stress by treating muscles with Tunicamycin (4 hours) and saw no effect on the integrity of the tubular lysosomal network. We confirmed the induction of ER stress by extracting protein from the same preps that were imaged and analyzing Hsc-70/BiP (a protein known to be induced by ER stress) levels by western blot. These data are included in new Figure 3.

https://doi.org/10.7554/eLife.07366.020

Article and author information

Author details

  1. Alyssa E Johnson

    Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    AEJ, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    No competing interests declared.
  2. Huidy Shu

    Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    HS, Conception and design, Acquisition of data, Analysis and interpretation of data
    Competing interests
    No competing interests declared.
  3. Anna G Hauswirth

    Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    AGH, Acquisition of data, Analysis and interpretation of data
    Competing interests
    No competing interests declared.
  4. Amy Tong

    Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    AT, Contributed unpublished essential data or reagents
    Competing interests
    No competing interests declared.
  5. Graeme W Davis

    Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    GWD, Conception and design, Drafting or revising the article
    For correspondence
    graeme.davis@ucsf.edu
    Competing interests
    GWD: Reviewing editor, eLife.

Funding

National Institutes of Health (R37NS047342)

  • Graeme W Davis

Jane Coffin Childs Memorial Fund for Medical Research (Postdoctoral Fellowship)

  • Alyssa E Johnson

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

Acknowledgements

We thank members of the Davis lab for helpful advice and critically reading the manuscript. We are grateful to Kevin Ford and Ryan Jones for providing custom scripts for EPSP and mEPSP analyses. We also thank TK Sang and B Ye for generously sharing fly stocks. AEJ is a fellow of the Jane Coffin Childs Memorial Fund for Medical Research. This study was supported by NIH grant NS047342 to GWD.

Reviewing Editor

  1. K VijayRaghavan, National Centre for Biological Sciences, Tata Institute for Fundamental Research, India

Publication history

  1. Received: March 7, 2015
  2. Accepted: July 11, 2015
  3. Accepted Manuscript published: July 13, 2015 (version 1)
  4. Version of Record published: August 14, 2015 (version 2)

Copyright

© 2015, Johnson 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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