When dissociated myotomal tissues from early Xenopus embryos were cultured on glass coverslips coated with a mixture of ECM proteins, containing entactin/nidogen, collagen and laminin (ECL), topologically complex aneural AChR clusters were observed mostly on the bottom surface of muscle cells in contact with ECL-coated substratum (Figure 1A). Some aneural AChR clusters could also be found within the top surface of muscle cells but exhibited a sparsely scattered morphology. To further identify if specific ECM proteins are required for the formation of these topologically complex AChR clusters, we tested several key ECM proteins individually, including laminin, collagen, and gelatin, for their ability to induce AChR cluster formation in cultured Xenopus muscle cells. We found that the formation of bottom AChR clusters could be effectively induced by all ECM proteins tested, in contrast to the negative control using poly-D-lysine (PDL), a polypeptide commonly used to promote cell attachment (Figure 1B). In immortalized C2C12 myotubes, aneural AChR clusters can undergo a topological transformation from a plaque to a perforated pretzel-shaped, and eventually to C-shaped arrays (Kummer et al., 2004). Hence, we further classified the aneural AChR clusters in Xenopus muscle cultures into scattered, plaque, perforated, and C-shaped based on their morphological features. A majority of ECM-induced aneural AChR clusters exhibited perforated structures. While the percentage of muscle cells with AChR clusters was largely reduced on PDL-coated substrate, the largest share of AChR clusters exhibited scattered structures (9.59% out of 13.33% total) (Figure 1C), similar to those structures found on the top muscle surface. As opposed to a previous study showing the topological transformation of AChR clusters in C2C12 myotubes (Kummer et al., 2004), most AChR clusters, regardless of their topological features, were gradually dispersed over 6 days in cultured Xenopus muscle cells (Figure 1—figure supplement 1).

Our previous study indicated that actin depolymerizing factor (ADF)/cofilin, one of the molecular components in PLSs, regulates the formation and maintenance of AChR clusters via vesicular trafficking (Lee et al., 2009). Here, we further performed immunostaining of several typical core markers (composed of F-actin and actin-associated proteins, such as ADF/cofilin, Arp2/3 complex, and cortactin) and cortex markers (composed of adhesion molecules, such as vinculin, talin, and paxillin) of PLSs. All these markers were found to be enriched in the perforations (arrows) of AChR clusters in Xenopus muscle cultures (Figure 1—figure supplement 2). As cortactin is highly enriched and involved in both early and late stages of PLS formation (Webb et al., 2006), we then used cortactin as an indicator of PLS localization in AChR clusters. PLSs were spatially concentrated in a majority of perforated AChR clusters induced by different ECM proteins (Figure 1D and E). In contrast, PLSs were not localized in AChR clusters, even in a small fraction of AChR clusters (2.16% out of 13.33% total) exhibiting perforated structures, on PDL substrates. These data suggest that different ECM proteins can induce the assembly of PLSs, which may in turn promote the formation of topologically complex structures of AChR clusters.

The data above showed that perforated AChR clusters can be effectively induced by gelatin (Figure 1B and C), an ECM protein that was previously shown to induce simple AChR plaques only (Kummer et al., 2004). This novel finding prompted us to adopt the well-established fluorescent gelatin degradation assay (Starnes et al., 2011) to investigate the possible association between matrix degradation and PLS localization in AChR clusters. After plating the muscle cells on FITC-gelatin-coated substratum, confocal microscopy was performed to visualize the extent of gelatin degradation in 3 day old live Xenopus primary muscle cells. We observed FITC-gelatin that was extensively degraded in close association with the perforated regions of aneural AChR clusters (Figure 2A). Next, we quantitatively measured the fluorescence intensity of FITC-gelatin in different AChR clusters and identified a linear relationship between the extent of gelatin degradation and the perforation area of aneural AChR clusters (Figure 2B). A similar spatially restricted gelatin degradation pattern was observed in cultured C2C12 myotubes (Figure 2C). Besides gelatin, immunostaining experiments also showed the spatial degradation of laminin at the perforated regions of AChR clusters in Xenopus primary muscle cells (Figure 2D).

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To examine if the focal degradation pattern of ECM proteins is mediated by MMP activity, we first used two broad-spectrum MMP inhibitors, BB-94 and Marimastat (BB-2516), which are known to be highly potent against MMP-1, -2, -3, -7, -9 and MT1-MMP (Cathcart et al., 2015). Although perforated AChR clusters could be identified in muscle cells treated with 5 µM BB-94 or 10 µM BB-2516, the degradation of fluorescent gelatin associated with these perforated AChR clusters was completely abolished (Figure 2E and F). Quantification further demonstrated that the initial formation of aneural AChR clusters in muscle cells was largely unaffected in the presence of MMP inhibitors (Figure 2G). To investigate if PLS-associated MMP activity regulates the topological remodeling of AChR clusters, we monitored the same group of muscle cells over 2 days after aneural AChR clusters had been formed. We observed a significant reduction in the number of aneural AChR clusters in control muscle cells, in contrast to BB-94 or BB-2516 treatment that caused a slight increase in the number of aneural AChR clusters (Figure 2H). To further explore if MMP activity regulates the topological remodeling of AChR clusters, we performed time-lapse imaging to monitor the dynamic changes in AChR cluster morphology and intensity in response to BB-94 treatment. AChR clusters were first labeled with fluorescent α-bungarotoxin (0 hr), then they were monitored for 96 hr after treatment. In control untreated cells, we observed a gradual dispersal of aneural AChR clusters over the 96 hr imaging period; however, such spontaneous dispersal and topological remodeling of AChR clusters were largely inhibited by BB-94 treatment within the same imaging period (Figure 2I and J). These data demonstrated that MMP activity is involved in the topological remodeling and dispersal of aneural AChR clusters.

The spatially restricted patterns of gelatin degradation suggested the possible involvement of membrane-type MMPs (MT-MMPs) in modulating ECM environment focally to regulate AChR cluster remodeling and dispersal. To test this, we first performed immunostaining of Xenopus muscle cells and detected endogenous MT1-MMP that was enriched in the perforations of aneural AChR clusters (Figure 3A). Next, we over-expressed MT1-MMP-mCherry construct in muscle cells via Xenopus embryo microinjection. In MT1-MMP-mCherry-overexpressing muscle cells, we observed extensive degradation of fluorescent gelatin in the entire cell area (Figure 3B), in contrast to the spatially restricted degradation pattern at the perforated regions of AChR clusters in control cells. In addition, MT1-MMP-mCherry overexpressing cells showed a lower percentage of ECM-induced bottom AChR clusters, but a higher percentage of top AChR clusters, in comparison to the wild-type control cells (Figure 3C). Quantitative analyses on those bottom AChR clusters in MT1-MMP-mCherry-overexpressing cells showed that the intensity of aneural AChR clusters was significantly reduced, which was accompanied by the extensive gelatin degradation (Figure 3D). Interestingly, BB-94 or BB-2516 treatment was able to partially revert the distribution of aneural AChR clusters from the top surface back to the bottom surface of MT1-MMP-mCherry-overexpressing muscle cells (Figure 3C). Consistent with that, the extensive degradation of fluorescent gelatin and the reduction of AChR intensity caused by MT1-MMP-mCherry overexpression were largely rescued by either BB-94 or BB-2516 treatment (Figure 3D). Our results indicated that the precise control of focal ECM degradation by MT1-MMP at the perforated sites plays an essential role in the formation of bottom aneural AChR clusters.

Figure 3

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To investigate the cytoskeletal involvement for intracellular trafficking of MT1-MMP, we next performed total internal reflection fluorescence (TIRF) microscopy to visualize the perimembrane fraction of GFP-tagged microtubule plus end-binding protein 1 (EB1-GFP) in live cultured muscle cells. Interestingly, we observed EB1-GFP comets to be highly enriched at the perforated aneural AChR clusters (Figure 4A). This pattern of EB1-GFP comets was not an artifact of overexpression, as a similar localization pattern of endogenous EB1 signals was also detected in fixed Xenopus muscle cells (Figure 4—figure supplement 1). By analyzing the density and trajectory of EB1-GFP comets in time-lapse images, we found that EB1-GFP comets were densely localized, but they showed lower mobility, at sites of aneural AChR clusters than other regions of the cell (Figure 4B and C). These data suggested that AChR clusters are the focal point of directed microtubule-based intracellular transport. Previous studies showed that cytoplasmic linker-associated protein (CLASP) and LL5β, a PLS cortex protein involved in NMJ development (Kishi et al., 2005), interact with microtubules to direct the vesicular trafficking of different proteins to the NMJ (Basu et al., 2015). Thus, we examined if CLASP is required for directing the movement of EB1-GFP comets to PLS-enriched AChR clusters using antisense morpholino oligonucleotide (MO)-mediated knockdown approach. As validated by western blot analysis, endogenous CLASP protein level was largely reduced in Xenopus embryos microinjected with CLASP-MO (Figure 4—figure supplement 2). In CLASP-MO muscle cells, TIRF microscopy showed the perimembrane signals of EB1-GFP that were primarily found at the center of perforations within aneural AChR clusters (Figure 4A). Although the density of EB1-GFP comets was only slightly reduced at AChR cluster region (Figure 4B), the average speed of EB1-GFP comets in the region of AChR clusters was significantly increased by 26.7% to 0.12 ± 0.004 μm/s in CLASP-MO muscle cells (Figure 4C), indicating that CLASP is involved in microtubule capturing at aneural AChR clusters.

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To further study the directed trafficking and local capturing of EB1-GFP at aneural AChR clusters, we performed TIRF in combination with fluorescence recovery after photobleaching (FRAP) experiments. Before photobleaching, EB1-GFP signals were primarily enriched at the edge of perforations of aneural AChR clusters in control cells but at the center of perforations in CLASP-MO muscle cells (Figure 4D). To better show the spatial patterns of EB1-GFP signals at the perforations of aneural AChR clusters in control versus CLASP-MO muscle cells, we have plotted the fluorescence intensity profiles of EB1-GFP and AChR across a perforated region of AChR clusters, as indicated in Figure 4D. In the control cell, the peaks of EB1-GFP intensity showed a slight lateral shift towards the center of a perforated region (Top chart, Figure 4E), indicating the preferential enrichment of EB1-GFP at the edge of perforations. In contrast, CLASP knockdown caused the peak of EB1-GFP intensity to cover a larger area of a perforated region, indicating that EB1-GFP is enriched relatively closer to the center of perforations (Bottom chart, Figure 4E). After photobleaching the region of perforated AChR clusters (dotted rectangles in Figure 4F), we performed time-lapse imaging that reflects the rapid recovery of EB1-GFP fluorescence signals (in seconds after photobleaching). Spatially, the recovery of EB1-GFP signals was detected near the edge of those perforations within AChR clusters in control cells (arrows in Figure 4F, Figure 4—video 1), but it was largely reduced in CLASP-MO muscle cells (arrowheads in Figure 4F, Figure 4—video 2), leading to a significant increase in the half-time of EB1-GFP signal recovery by 58% from 5.71 ± 0.83 s in control cells to 9.01 ± 0.71 s in CLASP-MO muscle cells (Figure 4G and H). Collectively, our data suggested that PLSs may direct the vesicular trafficking to AChR clusters through EB1-/CLASP-mediated microtubule-capturing mechanisms.

As microtubules serve as major tracks for vesicular trafficking in mammalian cells, the cortical microtubule organization plays a crucial role in the targeted delivery of secretory and membrane proteins via exocytosis (Noordstra and Akhmanova, 2017). To further investigate if MT1-MMP intracellular trafficking is mediated by microtubule-based transport, we performed dual-channel live-cell imaging on Xenopus muscle cells over-expressing both MT1-MMP-mCherry and EB1-GFP (Figure 5A). In region ‘i’, some MT1-MMP-mCherry vesicles (arrows) were found to be initially immobile at the beginning of this time-lapse series, until they were captured by a moving EB1-GFP comet (arrowhead), leading to a coordinated movement of both EB1-GFP and MT1-MMP-mCherry signals from 27 s to 29 s time-points in this imaging period (middle row, Figure 5A). Another example in region ‘ii’ indicated that MT1-MMP-mCherry vesicles (arrows) were able to move bidirectionally on microtubule structures, as visualized by the fading tracks (arrowheads) generated by moving EB1-GFP comets (bottom row, Figure 5A). The spatiotemporal correlation between MT1-MMP-mCherry and EB1-GFP signals was better appreciated by constructing kymographs using multiple timeframes along ‘K1’ and ‘K2’ lines (Figure 5B). In these two kymographs, most MT1-MMP-mCherry vesicles were found to be relatively immobile. Interestingly, we observed some displacement of MT1-MMP-mCherry vesicles immediately after EB1-GFP comets had come into contact with them (arrows in Figure 5B). Consistent with the subcellular localization of MT1-MMP-mCherry, immunostaining data revealed that endogenous MT1-MMP exhibited punctuated structures inside the muscle cells, and some of them were co-localized with vesicle-associated membrane protein 1 (VAMP1) (Figure 5—figure supplement 1). Taken together, these data illustrated that EB1 coordinates the vesicular trafficking of MT1-MMP in cultured muscle cells.

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To further visualize the directed trafficking and capturing of MT1-MMP vesicles at PLS-associated AChR clusters, we performed TIRF-FRAP experiments on MT1-MMP-mCherry-expressing muscle cells after AChR labeling. The cells with low expression level of MT1-MMP-mCherry were chosen, in which the formation of aneural AChR clusters was largely unaffected. After photobleaching, we detected local capturing of MT1-MMP-mCherry vesicles at the edge of perforations within aneural AChR clusters (white arrows, region ‘i’ in Figure 5C), as well as at the edge of AChR cluster periphery (red and green arrows, region ‘ii’ in Figure 5C). In kymographs constructed along ‘K3’ and ‘K4’ lines, we observed MT1-MMP-mCherry vesicles that were frequently captured at both perforations (arrowheads) and periphery (arrows) of AChR clusters over the 120 s imaging period after photobleaching (Figure 5D). As both sites are enriched with PLS cortex markers, vinculin and talin (arrowheads in Figure 1—figure supplement 2), we speculated that PLSs, through the cortex domains, can mediate site-directed capturing of MT1-MMP vesicles for their subsequent surface insertion.

As MT1-MMP-mCherry over-expression significantly inhibited the formation of AChR bottom clusters (Figure 3C), we further used MT1-MMP tagged with a pH-sensitive green fluorescent protein pHluorin (MT1-MMP-pHluorin) to verify if the surface expression of MT1-MMP produces a similar inhibitory effect on AChR clustering. In cultured muscle with low MT1-MMP-pHluorin expression level, MT1-MMP-pHluorin signals were spatially enriched at the perforated regions of aneural AChR clusters (arrow ) and caused only a slight inhibition on aneural AChR clustering (Figure 6A and B). At high MT1-MMP-pHluorin expression, the formation of AChR bottom clusters was significantly inhibited, leading to sparsely scattered appearance if observed. These data further confirmed that MT1-MMP activity is crucial for the formation of topologically complex AChR clusters by precisely controlling focal ECM degradation at the perforated sites.

To validate the surface localization of MT1-MMP in AChR clusters, we treated MT1-MMP-pHluorin-expressing muscle cells with a non-permeable buffer at pH 5.0 and observed a large reduction in MT1-MMP-pHluorin fluorescence intensity (arrows in Figure 6C), mimicking the quenching of MT1-MMP-pHluorin signals in acidic vesicular compartments. The signals were then restored after changing the culture medium back to pH 7.8, validating the effectiveness of this probe for visualizing surface localization of MT1-MMP, as it emits bright fluorescence upon surface insertion. Using this probe, we further determined the correlation between surface localization of MT1-MMP and topological remodeling of AChR clusters by monitoring the changes in the same AChR clusters over 24 hr in cultured muscle cells with different expression levels of MT1-MMP-pHluorin (Figure 6D). It should be noted that muscle cells with low MT1-MMP-pHluorin expression level were chosen to minimize the non-specific effects of exogenous overexpression. In this example, AChR cluster intensity was moderately reduced by 26% after 24 hr in the muscle cell with a very low expression level of MT1-MMP-pHluorin. In contrast, AChR cluster intensity in a muscle cell with a relatively higher (low) MT1-MMP-pHluorin expression showed about 78% reduction during the same period. Quantitative analyses on a pool data of 13 AChR clusters further indicated that MT1-MMP-pHluorin intensity in AChR cluster region was linearly correlated with the change in AChR cluster intensity over the next 24 hr (Figure 6E). These data indicated the functional role of surface MT1-MMP in promoting the topological remodeling of AChR clusters.

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Next, we performed TIRF-FRAP experiments that aimed to identify the exact locations of MT1-MMP surface insertion at AChR clusters. In this example, strong and discrete MT1-MMP-pHluorin signals re-appeared at the perforation (arrows) and periphery (arrowheads) of AChR clusters at time-points of 118 s and 48 s, respectively, in control muscle cells after photo-bleaching (Figure 6F). Kymographs constructed along ‘K1’ and ‘K2’ lines indicated that some newly inserted MT1-MMP-pHluorin signals were relatively stable (arrows), while some were short-lived but with multiple events of MT1-MMP surface insertion at the same location in AChR clusters (arrowheads) within a short imaging period (Figure 6G). By quantifying the number of events of MT1-MMP surface insertion, we detected a significant reduction of MT1-MMP-pHluorin surface insertion at AChR clusters in CLASP-MO muscle cells when compared to that in control cells (Figure 6H). Together with the above data showing the local capture of EB1-GFP comets, this data further confirmed that vesicular trafficking and surface targeting of MT1-MMP are spatially regulated by PLS cortex proteins at the perforation and periphery of AChR clusters through EB1-/CLASP-mediated microtubule capturing mechanisms.

To understand the functional role of PLS-associated MMP activity at developing NMJs, we first examined the effects of MMP inhibitors on ECM degradation and synaptic AChR clustering in Xenopus nerve-muscle co-cultures. Along the trail of neurites in contact with the basal membrane of muscle cells (arrows), we observed spatial degradation of fluorescent gelatin-coated substratum and synaptic AChR clustering at the nerve-muscle contacts (Figure 7A). Like the focal gelatin degradation patterns observed in AChR-poor regions of aneural clusters (Figure 2A), synaptic AChR clusters at the nerve-muscle contacts were found to be closely associated to, but not perfectly co-localized with, the sites of gelatin degradation, with only 17.15 ± 15% of area in AChR clusters to be associated with gelatin degradation in control co-cultures (Figure 7C). However, both gelatin degradation and nerve-induced AChR clustering were effectively inhibited by BB-94 or BB-2516 (Figure 7A and B). Nerve innervation of skeletal muscles involves local signals to initiate the formation of synaptic AChR clusters and global signals to induce the dispersal of aneural AChR clusters (Dai and Peng, 1998). Thus, we further examined the involvement of MMP activity in the dispersal of aneural AChR clusters induced by latex beads coated with agrin, a heparan sulfate proteoglycan that induces postsynaptic differentiation. Local application of agrin-coated beads to cultured muscle cells is capable of inducing postsynaptic differentiation in a spatiotemporally controllable manner (Lee et al., 2009). In control cells, we detected a gradual dispersal of aneural AChR clusters upon agrin-bead stimulation for 24 hr; however, those clusters in agrin bead-contacted muscle cells were greatly stabilized by BB-94 or BB-2516 (Figure 7D and Figure 7—figure supplement 1A). To determine if agrin stimulation enhances the localization of MT1-MMP in aneural AChR clusters prior to their dispersal, we performed immunostaining experiments, in which we demonstrated an increase of endogenous MT1-MMP signals at aneural AChR clusters upon agrin stimulation (Figure 7—figure supplement 2). These data implicated the important role of MT1-MMP activity in regulating not only the assembly of synaptic AChR clusters at nerve-muscle contacts, but also the disassembly of aneural AChR clusters, at developing NMJs.

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In contrast to nerve-induced AChR clustering, we noted that MMP inhibitors did not significantly affect the formation of AChR clusters induced by agrin beads (Figure 7—figure supplement 1B and C). As the latex beads were coated with a high amount of exogenous recombinant agrin, and they were placed on the top surface of cultured muscle cells, it remained unclear if MMP activity regulates synaptic AChR clustering induced by the physiological amount of agrin and in the presence of different ECM proteins. Therefore, we further developed an ‘endogenous agrin track’ assay (Figure 7E), by which we examined the effects of MMP inhibitors on synaptic AChR clustering in muscle cells cultured on ECL-coated substratum with endogenous agrin tracks. Using this assay, both BB-94 and BB-2516 were found to significantly inhibit the formation of synaptic AChR clusters induced by the endogenous agrin tracks (Figure 7F and G), consistent with our data on nerve-induced AChR clustering.

To identify the postsynaptic functions of MT1-MMP in NMJ development, antisense MO was used to knock down the endogenous expression of MT1-MMP specifically in muscle cells and then co-cultured with wild-type (WT) spinal neurons. The efficiency of MO-mediated knockdown of endogenous MT1-MMP protein level in Xenopus embryos was validated by western blot analysis (Figure 8—figure supplement 1). Pre-existing and newly inserted AChR molecules were differentially labeled by α-bungarotoxin conjugated with different fluorophores according to our previously established protocol (Lee et al., 2009). In short, we first labeled all pre-existing AChR molecules with tetramethylrhodamine-conjugated α-bungarotoxin in muscle cells. After plating the spinal neurons for 1 day, the newly synthesized AChRs can then be labeled with Alexa Fluor 488-conjugated α-bungarotoxin. In control co-cultures, both pre-existing and newly inserted AChR signals were detected at the nerve-muscle contact sites (Figure 8A), suggesting that both receptor pools contribute to the assembly of nerve-induced synaptic AChR clusters. Importantly, MT1-MMP knockdown caused a significant reduction, but not a complete inhibition, of nerve-induced AChR clustering in the chimeric co-cultures of MT1-MMP MO muscles with WT neurons (Figure 8A and B). This could be explained by the partial knockdown of endogenous MT1-MMP protein level (Figure 8—figure supplement 1) and/or the recruitment of diffuse AChRs to the nerve-muscle contacts via MT1-MMP-independent processes. Intriguingly, both pre-existing and newly inserted AChRs at the nerve-muscle contacts were equally affected in the chimeric co-cultures of MT1-MMP MO muscles with WT neurons (Figure 8C and D). To rule out the possible off-target effects of MT1-MMP MO, we next performed rescue experiments by overexpressing MT1-MMP-mCherry (MT1-mCherry) in MT1-MMP knockdown (MT1-MO) muscle cells. In the chimeric co-cultures of MT1-MO + MT1-mCherry muscles with WT neurons, we found that the percentage of nerve-muscle contacts with AChR clusters was largely returned to the level comparable with the control groups, and both pre-existing and newly inserted AChRs were detected at the nerve-muscle contacts (Figure 8). These data suggested that postsynaptic MT1-MMP regulates both the surface delivery of newly synthesized AChR molecules and the redistribution/recruitment of pre-existing AChR molecules (from either aneural AChR clusters or diffuse AChRs on muscle surface) during the assembly of nerve-induced postsynaptic specializations at developing NMJs.

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To determine if aneural AChR clusters contribute to the formation of NMJs, we next performed laser-based photobleaching experiments to differentiate the recruitment of AChR molecules from aneural clusters versus diffuse AChRs for the assembly of nerve-induced synaptic AChR clusters (Figure 9A). Without photobleaching, AChR signals were prominently detected at the nerve-muscle contact sites. The formation of nerve-induced synaptic AChR clusters was accompanied by the dispersal of aneural AChR clusters after 1 day in co-culture. In the experimental groups with photobleaching of aneural AChR clusters before co-culturing with the spinal neurons, we detected a significant reduction in AChR signals, which were contributed from diffuse AChRs only, at the nerve-muscle contacts in comparison to the control groups without photobleaching (Figure 9B and C). These results provided compelling evidence to support the hypothesis that AChR molecules from aneural clusters serve as a major source for the assembly of synaptic AChR clusters.

Figure 9

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To further investigate the involvement of MT1-MMP in the recruitment of AChRs from aneural to synaptic clusters, we performed the same photobleaching experiments using chimeric co-cultures composed of either control MO or MT1-MMP MO muscle cells with WT spinal neurons (Figure 9B). In the chimeric co-cultures with control MO muscle cells, both aneural clustered and diffuse AChRs were recruited to the nerve-muscle contacts at a level comparable to the wild-type co-cultures. However, knockdown of MT1-MMP in muscles significantly suppressed the recruitment of AChR molecules from aneural to synaptic clusters, while the recruitment from diffuse AChRs was unaffected (Figure 9C). Consistent with the data above (Figure 7D), we found that knockdown of muscle MT1-MMP caused a higher stability of aneural AChR clusters against nerve-induced dispersal (arrowheads, lower left panels in Figure 9B). Interestingly, after photobleaching of aneural AChR clusters in the chimeric co-cultures with MT1-MMP MO muscles cells, the fluorescence intensity of photobleached aneural AChR clusters was partially recovered after 1 day in co-culture (arrowheads, lower right panels in Figure 9B). These results suggested that diffuse AChRs were constantly recruited to aneural AChR clusters that were stabilized by muscle MT1-MMP knockdown, presumably through the diffusion-trap mechanisms in which diffuse AChRs can be immobilized by molecular scaffolds associated with the stable aneural AChR clusters (Geng et al., 2008). Taken together, our data further supported that muscle MT1-MMP plays a dual-functional role in regulating the formation and dispersal of synaptic and aneural AChR clusters, respectively.

To further investigate if MT1-MMP is required for NMJ development in vivo, we first performed immunostaining experiments to examine the localization of endogenous MT1-MMP at mature NMJs in rat soleus muscles. MT1-MMP was found to be highly enriched at the NMJs (Figure 10A). On the other hand, some immunostaining signals, which were partly contributed by intracellular MT1-MMP proteins, were also observed outside of the AChR cluster regions, further indicating that the precise control of MT1-MMP trafficking and surface insertion is required for mediating local ECM degradation at NMJs. The specificity of MT1-MMP antibody was validated by pre-incubating the antibody with recombinant MT1-MMP proteins, which caused a large reduction of immunostaining signals at the NMJs. To further determine whether MT1-MMP is postsynaptically localized, a denervation experiment on adult rat soleus muscle was performed (Figure 10B). In 4 days after sciatic nerve cut, we observed only a slight reduction in MT1-MMP signals at the NMJs of denervated muscles, suggesting that MT1-MMP is expressed primarily in postsynaptic muscles.

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Next, we examined the patterns of AChR clusters in embryonic diaphragm muscles between WT and MT1-MMP-deficient mice, which was previously established (Sakamoto and Seiki, 2009; Zhou et al., 2000). Specifically, diaphragms from WT and MT1-MMP^-/- mice at E13.5 and E18.5 were dissected and whole-mount stained for AChR and phrenic nerve. Aneural and synaptic AChR clusters were quantified based on whether the clusters were in direct contacts with the phrenic nerve branches. The endplate band width, which represents the zone of AChR cluster formation in the center of muscle fibers innervated by the phrenic nerve, was also measured. At E13.5, MT1-MMP^-/- mice showed a significantly higher density of aneural AChR clusters than the age-matched WT mice, but the density of synaptic AChR clusters was largely unchanged at this stage (Figure 10C-E). At E18.5, we observed a significant reduction in the density of synaptic AChR clusters and endplate band width in MT1-MMP^-/- mice compared to those in the WT mice (Figure 10E-F), indicating that synaptic AChR clusters formed in the knockout mice were immature with the altered number and density. Interestingly, the phrenic nerve in MT1-MMP^-/- mice also exhibited abnormal phenotype with less axonal defasciculation and arborization in the diaphragm muscles (Figure 10C and G), suggesting either the retrograde signaling of postsynaptic MT1-MMP or a possible role of neuronal MT1-MMP in regulating axonal growth and differentiation. Together, these findings further validated our in vitro data showing an important function of MT1-MMP in regulating the recruitment of aneural AChR clusters for the assembly of postsynaptic specializations at developing NMJs in vivo.

Adult Xenopus laevis animals were purchased from Xenopus 1. Xenopus oocytes were fertilized in vitro, and the embryos were raised in Holtfreter’s solution (vol/vol; 60 mM NaCl, 0.6 mM KCl, 0.9 mM CaCl₂, 0.2 mM NaHCO₃, pH 7.4) at 22°C. 20–100 pg of DNA constructs encoding MT1-MMP-pHluorin, MT1-MMP-mCherry (a gift from Dr Cheng-Han Yu, The University of Hong Kong) or EB1-GFP (Addgene, 39299) were microinjected into 1 cell stage Xenopus embryos using an oocyte injector, Nanoject (Drummond Scientific). GFP- or mCherry-expressing embryos were screened for primary culture preparation, as previously described (Lee et al., 2009; Peng et al., 1991). In short, dorsal parts of wild-type or microinjected embryos at Nieuwkoop and Faber stage 19–22 were dissected. After the enzymatic digestion by collagenase (Sigma, C98191G), myotomal tissues and neural tubes were isolated, followed by dissociation with calcium-magnesium-free solution. The dissociated cells were plated on either coverslips or glass-bottom dishes coated with a mixture of cell attachment substrate, entactin-collagen IV-laminin (ECL) (Millipore, 08–100), or specific ECM proteins including laminin (Sigma, L2020), collagen I (Fisher Scientific, C354249), and gelatin (Sigma, G1393) at 10 µg/ml, or poly-D-lysine (Sigma, P1024) at 100 µg/ml. Cells were maintained in culture medium containing 87% Steinberg’s solution (vol/vol; 60 mM NaCl (Sigma, S3014), 0.67 mM KCl (Sigma, P5405), 0.35 mM Ca(NO₃)₂ (Sigma, 202976), 0.83 mM MgSO₄ (Sigma, M2773), 10 mM HEPES (Sigma, H3375), pH 7.4), 10% Leibovitz's L-15 medium (Sigma, L4386) (vol/vol), 1% fetal bovine serum (Gibco, 10270) (vol/vol), 1% penicillin/streptomycin (Thermo Fisher Scientific, 15140122) (vol/vol) and 1% gentamicin sulfate (Thermo Fisher Scientific, 15750060) (vol/vol). Muscle cells were kept at 22°C for at least 24 hr to allow cell attachment and development of aneural AChR cluster before treatments, if any. To make nerve-muscle co-cultures, dissociated spinal neurons were plated into 2 d-old muscle cultures and grew for 1 day before imaging. Most of the nerve-contacted sites can be found in the basal membrane of muscle cells on ECM-coated substrates, where the growth cones of spinal neurons are able to crawl and migrate under the muscle cells (Anderson and Cohen, 1977a; Anderson et al., 1977b). All experiments involving Xenopus frogs and embryos were carried out in accordance with the approved protocol (#4627–18) by the Committee on the Use of Live Animals in Teaching and Research (CULATR) of The University of Hong Kong.

Authenticated C2C12 cell line was purchased from ATCC. Cell identity has been confirmed by STR profiling and cell line was found to be free of Mycoplasma. Cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) containing 20% fetal calf serum supplemented with penicillin/streptomycin. Only cells with low passage (<5) were used in this study. After trypsinization, cells were cultured on glass coverslips coated with 1 mg/ml fluorescent gelatin. To induce myoblast fusion, the culture medium was replaced with the fusion medium containing 2% horse serum (Thermo Fisher Scientific, 16050130) in DMEM supplemented with penicillin/streptomycin.

MT1-MMP^+/- mice in C57BL/6 background were obtained from Motoharu Seiki (Kanazawa University, Japan). In short, exons 1–5 encoding the catalytic domain of MT1-MMP were substituted by LacZ coding sequence (Sakamoto and Seiki, 2009). MT1-MMP^+/- mice were backcrossed against C57BL/6 background mice at least 12 times. MT1-MMP^+/- mice were mated to produce homozygous MT1-MMP^-/- mice, which were confirmed by genotyping. All animal experiments were carried out in accordance with the approved protocol (#5149–19) by the Committee on the Use of Live Animals in Teaching and Research (CULATR) of The University of Hong Kong.

6-week-old rats received anesthesia by intraperitoneal injection of 130 mg/kg ketamine with 13 mg/kg xylazine. After anesthesia, the left sciatic nerve was exposed with a lateral longitudinal straight incision from the greater trochanter to mid-thigh, and then by blunt dissection between the quadriceps femoris and biceps femoris. After clearing the surrounding connective tissues, the left sciatic nerve was cut with scissors. A sham operation was performed, where the sciatic nerve was exposed, and then the skin was closed immediately afterwards. After the surgery, the rats were returned to the cages in a pathogen-free environment, in a 12 hr light/dark cycle and with water and food ad libitum. Rats were euthanized 4 days after surgery with an overdose of pentobarbital (300 mg/kg). Then, soleus muscles were harvested from the control and denervated rats. After the fixation of the harvested muscles with 2% paraformaldehyde for 30 min, the tissues were then covered with tissue-tek O.C.T. compound (Sakura), and frozen with methanol in liquid nitrogen. Serial longitudinal and cross sections (20 μm thick) were cut in a cryostat microtome at −15°C (CryoStar NX50 Cryostat, Thermo Scientific). All animal experiments were carried out in accordance with the approved protocol by the Committee on the Use of Live Animals in Teaching and Research (CULATR) of The University of Hong Kong.

Knockdown of endogenous proteins in Xenopus was achieved by embryonic injection of antisense morpholino oligonucleotides (MO, Gene Tools), which bind to the target mRNA sequence to block its protein translation. The following MO sequences were used in this study: Xenopus MT1-MMP MO: 5′-CCAGG CTGCT CTCAG AGGCT CCATC-3′, Xenopus CLASP MO: 5’-GCCAG TAGTC CATTC CCTGT TCCAT-3’, and standard control MO: 5’-CCTCT TACCT CAGTT ACAAT TTATA-3’. To visualize the presence of MO in the microinjected embryos, MOs were co-injected with Alexa Fluor 488- or Alexa Fluor 546-conjugated dextran (Thermo Fisher Scientific, D22910 or D22911) as a cell lineage tracer. The effectiveness of MO-mediated knockdown of endogenous proteins was validated by Western blot analyses.

For the experiments studying the effect of MMP inhibitors on the formation of aneural AChR clusters, 5 μM BB-94 (ApexBio, A2577) or 10 μM Marimastat/BB-2516 (ApexBio, A4049) was added in the culture medium before muscle cell plating. For the experiments studying the nerve-induced or agrin bead-induced AChR clusters, BB-94 or BB-2516 was added to the muscle cultures about 1 hr before adding the spinal neurons or agrin beads.

Surface AChRs were labeled with 0.1 μM tetramethylrhodamine-, Alexa Fluor 488-, or Alexa Fluor 647-conjugated α-bungarotoxin (Thermo Fisher Scientific, T1175, B13422, or B35450) for 45 min in culture medium, followed by extensive washing with culture medium. To differentially label the pre-existing and newly-inserted pools of AChRs (Lee et al., 2009), the pre-existing AChRs were first labeled with tetramethylrhodamine-conjugated α-bungarotoxin for 45 min, followed by saturating all unlabeled surface AChRs with a high dose (6 μM) of unconjugated α-bungarotoxin (Thermo Fisher Scientific, B1601) for 30 min and then washed extensively with culture medium for at least 3 times. After 1 day, newly inserted AChRs were labeled with 1 μM Alexa Fluor 488-conjugated α-bungarotoxin. Glass coverslips with live cultured cells were then mounted on custom-made sealed chambers containing culture medium for live-cell imaging.

Glass coverslips (Fisher Scientific, 12-545-82) or glass-bottom dishes (MatTek, P35G-1.5–14 .C-GRID) were coated with 1 mg/ml Oregon Green 488-gelatin or FITC-gelatin (Thermo Fisher Scientific, G13186 or G13187) for 10 min and followed by cross-linking with 0.5% glutaraldehyde (Sigma, G5882) in PBS for 15 min. After that, gelatin-coated coverslips or glass-bottom dishes were treated with 5 mg/ml NaBH₄ (Sigma, 452882) for 3 min. Dissociated cells were plated on fluorescent gelatin-coated coverslips or glass-bottom dishes and were maintained for different time periods with or without treatment, as specified. All images were taken using the same acquisition settings across different experimental groups. The extent of gelatin degradation was quantified using ImageJ (NIH), by which the mean intensity of fluorescent gelatin signals within the regions of aneural AChR clusters or nerve-muscle contact sites was measured, and then normalized with the mean intensity at the adjacent region in the same cell (except in MT1-MMP-mCherry overexpression experiments, the mean intensity at the adjacent region without cell attachment was used for normalization).

Dissociated Xenopus spinal neurons were first plated on ECL-coated coverslips and cultured for 1 day. Spinal neurons were then digested and removed with 1% Triton X-100 in PBS for 5 min, followed by extensive washing with PBS for at least 5 times for 5 hr. For the experimental groups, pharmacological agents were added into the culture medium before plating the dissociated muscle cells. After 1 day in culture, the location of agrin tracks was visualized by agrin immunostaining (1:100; DSHB, 6D2) and then confirmed if neurites were not found in the phase contrast images.

Cultured cells were either fixed with 4% paraformaldehyde (Thermo Fisher Scientific, 28908) in PBS for 15 min, followed by permeabilization with 0.5% Triton X-100 in PBS for 10 min or fixed with −20°C 95% ethanol for 5 min. For EB1 immunostaining, muscle cells were fixed with 90% methanol containing 50 mM EGTA at −20°C for 15 min, followed by further fixation with 4% PFA at room temperature for 10 min. The fixed cells were washed extensively with PBS for at least 3 times, followed by blocking overnight with 2% Bovine Serum Albumin (Sigma, A9418) at 4°C. Cells were incubated with primary antibodies, including β-tubulin (1:1000; DSHB, 6G7-s),cortactin (1:500; Santa Cruz Biotechnology, sc11408), paxillin (1:100; Biolegend, 624001), p34-Arc (1:100; Millipore, 07–227), talin (1;100; Sigma, T3287), vinculin (1:100; Sigma, V4504), laminin (1:100; Thermo Fisher Scientific, PA516287), EB1 (1:100; BD Biosciences, 610534), Synaptobrevin/VAMP1 (1:300; Synaptic Systems,104002), MT1-MMP (1:50; Millipore, MAB3328), or ADF/cofilin (1:500; a gift from Dr James Bamburg, Colorado State University) at room temperature for 2 hr, followed by fluorophore-conjugated secondary antibodies (Thermo Fisher Scientific) for 45 min. For MT1-MMP immunostaining, muscle cells were fixed with −20°C 100% methanol for 5 min. Alexa Fluor 488 Tyramide SuperBoost kit (Thermo Fisher Scientific, B40912) was also used. In short, endogenous peroxidase activity was first quenched by incubating the fixed muscle cells with 3% hydrogen peroxide for 1 hr, followed by blocking with 2% Bovine Serum Albumin for 1 hr at room temperature. The fixed muscle cells were then stained with MT1-MMP primary antibody at 4°C overnight, followed by incubating with poly-HRP-conjugated secondary antibody for 1 hr at room temperature. Endogenous MT1-MMP signals were developed by incubating the cells with Alexa Fluor 488 tyramide solution for 5 min. Coverslips were then mounted on glass slides with the anti-bleaching reagent fluoromount-G (Thermo Fisher Scientific, 00-4958-02) for later observation.

For mouse diaphragm muscle tissues, the entire diaphragms were dissected and fixed in 2% paraformaldehyde (PFA) in PBS at room temperature for 1 hr, and then incubated with 0.1 M glycine in PBS for 30 min. The fixed tissues were labeled with 0.1 μM tetramethylrhodamine α-bungarotoxin at room temperature for 30 min. After extensive washing with PBS, tissues were labeled with the primary antibody against neurofilament NF200 (1:1000; Sigma, N4142) in the blocking buffer (1% BSA and 0.3% Triton X-100) at 4°C for 2 days. After extensive washing with 0.5% Triton X-100 in PBS, tissues were incubated overnight with Alexa Fluor 488-conjugated secondary antibody at 4°C. Diaphragm muscles were flat-mounted with fluoromount-G onto glass slides.

For rat soleus muscle tissues, the sections were incubated with 0.1 M glycine for 30 min to remove autofluorescence background and permeabilized by methanol. After blocking with 2% BSA and 0.2% Triton X-100 at room temperature for 2 hr, the sections were labeled with primary antibody against MT1-MMP (1:100; Santa Cruz Biotechnology, sc-373908 and 1:100; Millipore, MAB3328), neurofilament NF200 (1:500; Sigma, N4142), and synaptophysin (1:1000; Abcam, ab32127) for 2 days at 4°C. For MT1-MMP pre-incubation, 1 μg recombinant human MT1-MMP (R and D Systems, 918-MP-010) was pre-incubated with 1 μg MT1-MMP antibody for 2 hr at room temperature. After extensive washing with PBS, the muscle sections were labeled with fluorophore-conjugated secondary antibodies (Thermo Fisher Scientific) for 2 hr at room temperature. Muscle sections were mounted with DAPI-Fluoromount-G (Electron Microscopy Sciences, 17984–24).

Xenopus embryos at Nieuwkoop and Faber stage 19–22 were homogenized in RIPA buffer in the presence of protease inhibitor cocktail and EDTA, followed by incubation on ice for 5 min. After high-speed centrifugation (15000 x g), the supernatant was collected for protein concentration measurement using a BCA protein assay kit (Thermo Fisher Scientific, 23227). 30 μg protein lysates were used for protein separation by SDS-PAGE, and then blotted onto Immobilon-P membrane (Millipore, IPVH00010). After blocking with 5% non-fat milk in TBST, the blots were probed for the following primary antibodies: MT1-MMP catalytic domain (1:1000; Millipore, AB6005; kindly provided by Professor Timothy Gomez, University of Wisconsin-Madison); CLASP1 (1:250; Abcam, Ab85919); anti-β actin (1:1000; Sigma, A2228); anti-α tubulin (1:2000; Sigma, T6074) overnight at 4°C. After extensive washing with TBST, the blots were reacted with secondary antibodies conjugated with HRP. Signals were detected by ECL western blotting substrate (Thermo Fisher Scientific, 32106), and images were taken by ChemiDoc XRS+ System (Bio-Rad).

Wide-field fluorescence imaging was performed on inverted microscopes (IX81 or IX83, Olympus) using oil immersion PlanApo 60X NA 1.42 objective lens. On IX81, digital still images were captured by iXon EMCCD camera (Andor) through the software ) through the software cell^R (Olympus). On IX83, digital still and time-lapse images were captured by ORCA-Flash4.0 LT+ digital CMOS camera (Hamamatsu) through the software MicroManager (Open Imaging) (Edelstein et al., 2014).

In FRAP experiments, total internal reflection fluorescence (TIRF) mode using Axio TIRF unit fitted in an inverted microscope equipped with oil immersion 100X NA 1.46 DIC objective lens was used. Images were captured through Metamorph (Molecular Devices) by Evolve 512 EMCCD camera (Photometrics). To obtain the baseline of fluorescence intensity before photobleaching, 5 to 6 images were taken with 3 s intervals on cells expressing EB1-GFP and 1 s intervals on cells expressing MT1-MMP-pHluorin or MT1-MMP-mCherry. Photobleaching was performed by using Sapphire laser line (488 nm) with 50% laser intensity, which was kept the same for all FRAP experiments. The photobleaching duration was adjusted based on the signal intensity, ranging from 0.13 to 4.5 s for photobleaching areas between ~ 27 and 34 μm². After photobleaching, images were taken every 3 s until the fluorescence intensity has reached the plateau level.

In the experimental groups involving photobleaching of aneural AChR clusters, interactive bleaching function was used in a confocal microscope (LSM 800, Carl Zeiss) using 20X NA 1.4 DIC objective lens. Photobleaching was performed by using an argon laser (488 nm) with a 100% laser power. Identical settings were applied in all photobleaching experiments. A fluorescence image was taken immediately after photobleaching to ensure the signals of aneural AChR clusters had been completely bleached.

For imaging of mouse diaphragm muscles and rat soleus muscles, multiple tile-scanned, z-stack images were captured on a confocal microscope (LSM 800, Carl Zeiss) using 20X NA 1.4 DIC objective lens or oil immersion 63X NA 1.4 DIC objective lens. Images were captured with 32-channel GaAsp photomultiplier modules through the software ZEN 2.3 (Carl Zeiss). All acquisition settings (i.e. laser intensity and gain) were kept the same for different experimental groups in the same experiment. All acquired images were processed and analyzed by ImageJ (NIH) or Imaris (Bitplane).

To quantify the percentage of AChR top and bottom clusters, muscle cells were first identified in the phase-contrast channel, and then the presence of top or bottom aneural AChR clusters was scored in the fluorescence channel.

To quantify the extent of fluorescent gelatin degradation, the intensity of fluorescent gelatin in the region of interest (ROI, created based on the shape and location of AChR clusters) was measured and then normalized with the intensity of fluorescent gelatin at the adjacent area in the same muscle cell. For experiments involving MT1-MMP-mCherry overexpression, the fluorescent gelatin intensity in the region of AChR clusters was normalized with the intensity in the region without cell attachment.

To quantify the change in the area of aneural AChR clusters, ROI was created at the region of AChR clusters in the image taken at the first time point, and then applied to the images taken at subsequent time points for measuring and calculating the percentage change in cluster size.

To quantify the normalized intensity of aneural AChR clusters, the fluorescence intensity of AChRs in the ROI was first measured in all experimental groups and then normalized to the average AChR intensity in control group of the same experiment.

To quantify the density and speed of EB1 comets, ROI for AChR region was created based on the shape and location of AChR clusters, while ROI for non-AChR region was also created at the adjacent area in the same muscle cell. The number and speed of EB1 comets were measured from multiple frames in 7 s time-lapse images using TrackMate plugin in ImageJ (Tinevez et al., 2017). The density of EB1 comets was calculated by dividing the number of EB1 comets by the ROI area.

To quantify the FRAP experiments, the relative fluorescence intensity in the photobleached area was first measured and then normalized by the mean fluorescence intensity of the same region in the images acquired before photobleaching. The recovery half-time was determined by the time required for 50% recovery of fluorescence signals in the FRAP regions.

To quantify the number of MT1-MMP-pHluorin surface insertion events, ROI was created at the region of aneural AChR clusters. After photobleaching, the number of MT1-MMP-pHluorin signals in the ROI was measured from multiple frames in 60 s time-lapse images using TrackMate plugin in ImageJ. It was then normalized to the area of aneural AChR clusters.

To quantify the percentage of nerve-muscle contacts with AChR clustering, muscle cells with nerve contact at the bottom surface were first identified in the phase-contrast images, and then the presence of AChR clusters was scored in the fluorescence images.

To quantify the fluorescence intensity of nerve-induced AChR clusters per unit length of nerve-muscle contact, the integrated fluorescence value of AChR signals was first measured, and then it was normalized to the area of nerve-muscle contacts as determined and measured in the phase-contrast images.

To quantify the density of aneural and nerve-induced synaptic AChR clusters in mouse diaphragm muscles, the total number of AChR clusters was first measured from stack images using Imaris software. The number of nerve-induced synaptic AChR clusters was calculated based on the overlapping of signals between AChR clusters and neurofilaments. The number of aneural AChR cluster was then calculated by subtracting the number of nerve-induced synaptic AChR clusters from the total number of AChR clusters. The calculated numbers were then normalized to the length of the main nerve trunk. Endplate band width was defined by the average distance between two farthest AChR clusters in the muscle fibers along the main nerve trunk. The length of nerve branches was quantified by measuring the average axonal length from the main nerve trunk. Approximately 100 measurements were taken in each diaphragm, and the quantification of the endplate band width and length of branches was performed using the whole diaphragm from at least 3 animals in each genotype.

In all figures, mean and SEM values were shown in the graphs, unless otherwise specified. The numbers of biological replications and the statistical tests applied were specified in the figure legends.

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

This project was partly supported by the Early Career Grant (27102316) and General Research Fund (17100718 and 17100219) from Research Grants Council of Hong Kong, the Health and Medical Research Fund (04151086) from Food and Health Bureau of Hong Kong, and the Seed Fund Programme for Basic Research from The University of Hong Kong (201601159003, 201711159098, and 201811159078) to CWL. HRK is partly supported by HKU Postgraduate Fellowships in Integrative Medicine. We thank Dr Weichun Lin (UT Southwestern) for sharing the whole-mount diaphragm staining protocol, and Dr Timothy Gomez (University of Wisconsin-Madison) for providing the MT1-MMP antibody.

Animal experimentation: All experiments involving research animals were carried out in accordance with the approved protocols (#4627-18 and #5149-19) by the Committee on the Use of Live Animals in Teaching and Research (CULATR) of The University of Hong Kong.

1. Received: December 12, 2019
2. Accepted: February 10, 2020
3. Version of Record published: March 24, 2020 (version 1)

© 2020, Chan et al.

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