We previously demonstrated that kinesin-1 slides microtubules against each other, and this sliding generates the forces that drives outgrowth at the initial stages of neurite outgrowth (Lu et al., 2013b) and axon regeneration (Lu et al., 2015). Because kinesin-1 is a plus-end microtubule motor, it can only slide microtubules with their minus-ends leading and plus-ends trailing (Figure 1A). If this model is correct, it suggests that kinesin-1 must extend neurites by pushing microtubule minus-ends against the plasma membrane during the initial stages of neurite formation. Furthermore, because the model predicts that two microtubules have to be in antiparallel orientation to slide against each other, sliding by kinesin-1 will result in the simultaneous transport of two microtubules in opposite directions (see Figure 1A and the legend for the explanation). Bidirectional microtubule movement can indeed be observed in growing axons of cultured Drosophila neurons using tubulin tagged with a photoconvertible probe (Video 1).

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To initially test this hypothesis, we first took advantage of Drosophila S2 tissue culture cells. S2 cells provide a good model system to explore the mechanism of process formation because they canbe induced to form cellular processes when the integrity of the actin filament network is impaired by treatment with either Cytochalasin D or Latrunculin B (LatB) (Kim et al., 2007; Lu et al., 2013a). In addition, this system enables us to efficiently study the mechanisms of process formation by knocking down candidate proteins with double-stranded RNA (dsRNA) (Rogers and Rogers, 2008).

To study microtubule minus-end distribution in live cells, we ectopically expressed a fluorescently tagged microtubule minus-end binding protein called calmodulin-regulated spectrin-associated protein (CAMSAP), also known as Patronin or Nezha. CAMSAP proteins bind to microtubule minus-ends and stabilize them against depolymerization, making them the perfect candidate to label microtubule minus-ends (Akhmanova and Hoogenraad, 2015). We initially performed experiments with GFP-tagged Patronin, the single Drosophila member of CAMSAP family (Wang et al., 2013), but its expression level in S2 cells was very low and GFP signal was not robustly found on microtubules (data no shown). On the other hand, its mammalian ortholog CAMSAP3 tagged with GFP expressed at consistently higher levels and reliably decorated microtubule ends (Figure 1B).

First, we wanted to test whether GFP-CAMSAP3 decorates only one end of microtubules in Drosophila cells. Because the microtubule network is normally too dense to identify both ends of microtubules, we induced the formation of short microtubules in cells by partial depolymerization with 25 µM Vinblastine for 1 hr. Examination of these short microtubule fragments demonstrated that only one end of each microtubule contained a GFP-CAMSAP3 patch (Figure 1—figure supplement 1A). In untreated S2 cells, spontaneous growth and shrinkage events associated with the dynamic instability of microtubule plus-ends were not seen in microtubule ends decorated by GFP-CAMSAP3, suggesting that GFP-CAMSAP3 labels microtubule minus-ends in Drosophila cells (Video 2). Furthermore, EB1-GFP and mCherry-CAMSAP3 never colocalized when expressed in the same cell, further confirming the minus-end localization of CAMSAP3 (Figure 1—figure supplement 1B and 1C; Video 3). These results, together with published data (Tanaka et al., 2012; Hendershott and Vale, 2014; Jiang et al., 2014; Akhmanova and Hoogenraad, 2015), demonstrated that mammalian GFP-CAMSAP3 reliably marks microtubule minus-ends in Drosophila.

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To study localization of microtubule minus-ends in growing processes, we induced the formation of processes in S2 cells expressing GFP-CAMSAP3 and started collecting images 5 min after plating the cells. At this time point, nascent processes were actively growing. We simultaneously tracked GFP-CAMSAP3 and the plasma membrane using a membrane dye (CellMask Deep Red). We found that a significant fraction of growing processes contained GFP-CAMSAP3 dots at their tips, and that these processes only elongated when microtubule minus-ends were present at their tips (Figure 1C,D; Video 4). Interestingly, we often observed retraction of the GFP-CAMSAP3 marker from the process tip; these events always coincided with a pause in process outgrowth (Figure 1C, inset). Quantitative analysis demonstrated that while CAMSAP3 almost always colocalized with the tips of the growing processes, the plus-ends marker EB1 could only be found in the tips of the growing processes approximately 30% of the time (Figure 1E,F), suggesting that at this stage microtubule dynamics does not play a major role in process outgrowth.

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To investigate the localization of microtubule minus-ends in Drosophila neurons, we created a transgenic fly that expresses GFP-CAMSAP3 under the UAS promoter. Neurons were harvested and cultured from the brains of larvae expressing GFP-CAMSAP3 driven by the pan-neuronal promoter elav-Gal4 (Egger et al., 2013; Lu et al., 2015) (see Material and methods). We visualized microtubule minus-ends in growing neurites at the initial stages of growth (4 hr after plating), when neurons started to develop processes (length= 9.96 µm, s.d. ± 4.5 µm, n=50 axons). We found that, like in S2 cells, the growing neurites contained GFP-CAMSAP3 dots at the tips of neurites, and localization of the dots to the tips precisely correlated with neurite outgrowth (Figure 1G–I; Video 5). All together, these data show that at least a fraction of microtubules in growing neurites have the ‘wrong’ orientation (minus-end-out). Localization of microtubule minus-end(s) at the neurite tip correlates with neurite outgrowth, consistent with kinesin-1 pushing the minus-ends of microtubules against the plasma membrane driving the initial outgrowth.

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If kinesin-1 slides antiparallel microtubules at the initial stages of axon formation, the growing neurites should initially contain microtubules with mixed orientation (Figure 1A and the legend for the explanation). To test this prediction, we imaged and tracked the direction of the plus-end microtubule marker, EB1-GFP, in the axons of cultured neurons at different time points after plating. Quantitative analysis of EB1-GFP comets using tracking software (see Material and methods) demonstrated that, shortly after plating, growing neurites contained EB1 comets moving in both anterograde and retrograde directions (Figure 2A,B; Video 6). The microtubule orientation in axons remained mixed during the 1st day in culture. At 36 hr, the fraction of retrograde EB1 comets started to decline, and at 48 hr, developed axons were mostly filled with plus-end-out microtubules (Figure 2A,B; Video 6). These results demonstrated that axons initially contain microtubule arrays with mixed orientation.

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Our data suggest that some sorting mechanism is responsible for the elimination of microtubules with the ‘wrong’ (plus-ends toward the soma) orientation from maturing axons. We hypothesized that this sorting factor is cytoplasmic dynein because it has been reported that mutations in dynein light intermediate chain or dynein-cofactors (NudE) are required for uniform microtubule orientation in axons of Drosophila dendritic arborization (da) neurons (Zheng et al., 2008; Arthur et al., 2015). To characterize the role of dynein, we expressed two different shRNAs targeting dynein heavy chain (DHC) using elav-Gal4 to specifically knock down dynein in neurons. elav>DHC RNAi animals developed to the third instar larval stage; however, their locomotion was severely impaired and most died before reaching the pupae stage (data not shown). In addition, none of the surviving elav>DHC RNAi pupae eclosed into adults. We cultured neurons obtained from brains of third instar elav>DHC RNAi larvae. Our lab has previously shown that mitochondrial movement was substantially diminished in elav>DHC RNAi neurons, indicating that the activity of dynein is impaired in those neurons (Lu et al., 2015).

To track the microtubule orientation after dynein depletion, we genetically combined transgenes encoding DHC RNAi with EB1-GFP or GFP-CAMSAP3. We first quantified the direction of the EB1 comets in neurons grown for 48 hr; at this time point, microtubules in the axons of control neurons are mostly oriented with plus-end-out (Figure 2B). Analysis of EB1 comets in elav>DHC RNAi 48 hr-neurons revealed that axons contained microtubule arrays of mixed orientation (Figure 2C,D; Video 7), suggesting that dynein is necessary to remove minus-end-out microtubules from axons. Interestingly, while microtubule minus-ends in control neurons, as visualized by GFP-CAMSAP3, were scattered throughout the length of the axons, inactivation of dynein resulted in dramatic accumulation of the minus-end markers at neurite tips (Figure 2E; control, left panels; DHC-RNAi, right panels).

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To further investigate the role of dynein in organizing microtubule arrays in cell processes, we again examined Drosophila S2 cells. The treatment of S2 cells with DHC dsRNA efficiently knocked down DHC (Figure 2—figure supplement 1A,B). Recapitulating the neuronal phenotype, dynein depletion in S2 cells resulted in processes with EB1-GFP comets traveling in both directions (Figure 2F,G; Video 8). We also examined the distribution of the GFP-CAMSAP3 in S2 processes. In control cells, GFP-CAMSAP3 has a broad, scattered dot distribution (Figure 2H, left panel). Dynein depletion induced a striking en masse accumulation of CAMSAP3 at the tips of processes (Figure 2H,I). Overexposed images of those processes revealed that minus-ends could be still found in other areas of the processes (Figure 2—figure supplement 2). Identical phenotypes (mixed polarity of GFP-EB1 comets and accumulation of CAMSAP3 at the tips of the processes) were observed after knocking down several dynein cofactors (p150^Glued, Lis1 or NudE; Figure 2G–I; Figure 2—figure supplement 1B for knockdown efficiency). The en masse accumulation of microtubule minus-ends at neurite tips and S2 processes in dynein RNAi cells is consistent with the idea while kinesin-driven sliding stays intact, the sorting mechanism is inactivated by dynein depletion.

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In many organisms, the organization and orientation of the mitotic spindle requires attachment of cytoplasmic dynein to the cellular cortex (Laan et al., 2012; Kiyomitsu and Cheeseman, 2013). We hypothesized that organization of microtubules in axons, like their organization in the mitotic spindle, requires anchoring of dynein to the cortical network of actin filaments. To test this idea, we depolymerized actin filaments in cultured Drosophila neurons by treatment with LatB and quantified microtubule orientation in axons. We found that increasing concentrations of LatB gradually reduced the amount of F-actin, as judged by the intensity of rhodamine-phalloidin staining (data not shown). In parallel with F-actin depolymerization, axons displayed an increased fraction of microtubules with plus-ends facing the cell body (Figure 3A). At a high concentration of LatB (10 µM), when virtually no F-actin can be detected, about half of all EB1 comets moved toward the cell body, demonstrating that depolymerization of actin induced random microtubule orientation in neurons (Figure 3A). Treatment with high LatB concentration also induced accumulation of microtubule minus-ends at the tips of the axons in elav>GFP-CAMSAP3 neurons (Figure 3B).

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These data show that actin depolymerization phenocopied dynein depletion in axons, suggesting that cortical dynein sorts microtubules. If this hypothesis is correct, LatB washout should restore dynein recruitment to the cortex, and the microtubule sorting activity of cortical dynein should remove minus-ends-out microtubules from the tip of the axon. To test this prediction, we washed out LatB and tracked the localization of GFP-CAMSAP3. We observed that the LatB washout induced removal of GFP-CAMSAP3 from axon tips without affecting the length of the axon (Figure 3B). It should be mentioned that LatB washout did not result in the normal scattered distribution of minus-ends along the length of the axon observed in control neurons (Figure 2E, left panels). Instead, they remained clustered in the shaft, most likely because by the time of drug washout these minus-end-out microtubules were already crosslinked into a bundle.

To observe the sorting activity of cortical dynein in real time, we again used S2 cells. As in the case of neurons, the treatment of S2 cells with high concentration of LatB induced formation of processes containing microtubules of mixed polarity (Figure 3C) and massive accumulation of minus-ends in their tips (Figure 3D–F; Video 9). Strikingly, LatB washout resulted in robust movement of GFP-CAMSAP3 labeled minus-ends from the tips of processes toward the cell body (Figure 3F, middle panel; Figure 3G; Video 9). However, if the same experiment was performed after dynein knockdown, retrograde transport of microtubule minus-ends was not observed and the minus-ends caps stay at the tips of processes (Figure 3F, right panel; Video 9), directly demonstrating the role of cortical dynein in removal of minus-end-out microtubules from processes to the cell body.

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To further confirm that dynein and actin filaments are components of the same microtubule-sorting pathway, we decided to bypass the requirement for actin filaments by directly recruiting dynein to the cell membrane. For these experiments, we used a dynein recruitment tool developed by the Akhmanova and Hoogenraad labs (Hoogenraad et al., 2003; Kapitein et al., 2010), which contains the dynein activator Bicaudal D (BicD) fused to a FRB domain (FRB-GFP-BicD). The FRB-BicD-dynein complex can then be recruited to any region of interest by coexpression of a targeting protein coupled to a FKBP domain. This FKBP domain chemically dimerizes with the FRB domain in the presence of rapalog (a cell-permeable small molecule analog of rapamycin) (Clackson et al., 1998). To confirm that the FRB-FKBP dimerization system for dynein recruitment works in Drosophila S2 cells, we first coexpressed a GFP construct fused to FRB (FRB-GFP) and the peroxisome membrane-targeting signal peptide coupled to the red fluorescent protein (PEX3-RFP-FKBP). In the absence of rapalog, the GFP signal appeared soluble in the cytoplasm. The addition of rapalog recruited FRB-GFP to peroxisomes (Figure 3—figure supplement 1A). We next used this system to directly recruit dynein to the plasma membrane. For membrane targeting, we fused the transmembrane domain of GAP-43 (Heim and Griesbeck, 2004) with FKBP and coexpressed this construct with FRB-GFP-BicD in S2 cells. To visualize the recruitment of BicD to the membrane in these cells, we used total internal reflection fluorescence (TIRF) microscopy to image the GFP signal before and after addition of rapalog. Quantification showed that addition of rapalog significantly increased the intensity of the GFP fluorescence due to recruitment of cytoplasmic BicD to the plasma membrane (Figure 3—figure supplement 1B).

We next tested if direct recruitment of dynein to the membrane using rapalog could drive microtubule sorting in the absence of cortical actin. To test this hypothesis, mCherry-CAMSAP3 was used to track the localization of microtubule minus-ends. In the absence of rapalog, CAMSAP3 accumulated in the processes of S2 cells treated with 10 µM LatB. Time-lapse imaging before addition of the drug revealed that these CAMSAP3 clusters, marking positions of minus-ends, were either static or pushing against the plasma membrane (Video 10). However, when the BicD-dynein complex was recruited to the membrane by addition of rapalog, CAMSAP3-labeled microtubule minus-ends robustly moved away from the tip, toward the cell body (Figure 3H, top panels; Figure 3I; Video 10). The same assay performed in DHC knockdown cells did not show this removal of microtubule minus-ends (Figure 3H, bottom panels; Video 10). Taken together, these data confirmed that dynein is responsible for microtubule sorting, and this activity requires the attachment of the motor to the cortex mediated by actin filaments.

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We previously demonstrated that initial neurite formation in Drosophila neurons requires microtubule-microtubule sliding driven by kinesin-1. Knockdown of kinesin-1 in primary neurons impairs the motility of interphase microtubules and causes neurons to fail to develop axons (Lu et al., 2013b). This was surprising because kinesin-1 is a plus-end microtubule motor and thus can only slide microtubules with minus-ends leading and plus-ends trailing. Furthermore, symmetry considerations dictate that kinesin slides antiparallel microtubules against each other; if microtubules are oriented parallel to each other kinesin would bundle rather than slide them (Figure 1A). Both of these considerations appear to contradict established literature about microtubule polarity in axons.

In this work, we performed two experiments that further support the idea that kinesin-1’s sliding activity drives the initial stages of neurite formation (Lu et al., 2013b). First, nascent growing neurites contain microtubule arrays with mixed polarity. As mentioned above, this antiparallel orientation is required for microtubule sliding by kinesin-1 (Figure 1A). Second, microtubule minus-ends are pushed against the plasma membrane at the tips of processes, generating the force for neurite outgrowth.

Recent experimental data revealed that ‘mitotic’ kinesins, such as kinesin-5 (Nadar et al., 2008; Nadar et al., 2012) and kinesin-6 (Lin et al., 2012; del Castillo et al., 2015), play an important role in the regulation of axon outgrowth. The well established function of these mitotic motors is to reorganize and stabilize the mitotic spindle to facilitate chromosome segregation. These motors accumulate in the spindle midzone where antiparallel microtubules coming from opposite poles overlap. Our work shows that developing Drosophila axons, like the spindle midzone, are filled with antiparallel microtubule arrays. This microtubule configuration suggests that the same mechanisms controlling the mitotic spindle could be used to regulate axonal outgrowth (Baas, 1999). Interestingly, ‘mitotic kinesins’ control neurite outgrowth both in mammalian and Drosophila systems, which supports the idea that the molecular players that drive axon initiation and outgrowth in Drosophila may be conserved across species.

Our data show that minus-end-out microtubules are consistently observed at the early stages of neurite formation, while at later stages the vast majority of microtubules in axons have their plus-ends out, as have been reported in multiple published papers (Baas et al., 1988; Stepanova et al., 2003; Stone et al., 2008). Therefore, neurons must activate an additional sorting mechanism that eliminates these ‘wrong’ polarity microtubules from axons. Previous studies reported that dynein and dynein-associated proteins are required for correct microtubule orientation in axons of Drosophila da sensory neurons (Zheng et al., 2008; Arthur et al., 2015). Our results using cultured neurons are in full agreement with these studies, as dynein knockdown caused mixed microtubule polarity in axons. Interestingly, several studies have linked dynein-mediated microtubule reorganization to actin, both in interphase (Mazel et al., 2014) and in mitosis (Kotak et al., 2012; Kiyomitsu and Cheeseman, 2013). In this study, we showed that depolymerization of actin filaments (including cortical actin) using high concentrations of LatB causes the same defects in microtubule orientation as dynein knockdown. Recovery of cortical actin after drug washout results in efficient dynein-driven removal of minus-end-out microtubules from the tips of processes. Furthermore, the requirement for actin can be efficiently bypassed by direct recruitment of the dynein machinery directly to the plasma membrane. Therefore, we propose that microtubule sorting depends on the activity of cytoplasmic dynein attached to the cortical actin filament meshwork. However, as we did not get complete recovery of the wild type distribution of microtubules in our LatB washout or dynein-membrane recruitment experiments, it is likely that additional proteins that link actin and microtubule pathways in the neuron are involved in microtubule organization; one good candidate is the well characterized actin-microtubule crosslinking protein Short stop (Lee and Kolodziej, 2002).

In addition to the sorting activity described in this work, other groups have observed that dynein activity is required for axon elongation (Ahmad and Baas, 1995; Ahmad et al., 1998; Grabham et al., 2007). A more recent study by Miller et al. has reported that dynein generates forces that push the cytoskeletal meshwork forward during axonal elongation in cultured chick sensory neurons (Roossien et al., 2014). In agreement with their data, we predict that cortical dynein can promote axon outgrowth after the initial stages of neurite outgrowth, when microtubules are sorted into a plus-end-out orientation. Because the direction of dynein-powered forces is dictated by the intrinsic orientation of microtubules, microtubules with minus-end-out are redirected toward the cell body, whereas plus-end-out microtubules are transported toward the tip of the axon. Therefore, dynein-driven microtubule transport can not only remove microtubules with the wrong orientation (minus-end-out) but also push microtubules of the right orientation (plus-end-out) toward the axon tip.

In addition to the role of dynein in developing neurons, dynein may play an important role in axonal microtubule maintenance. Microtubule polymers, like other protein structures, require subunit turnover to maintain their integrity. It has been proposed that several microtubule severing proteins are able to fragment microtubules into short pieces (McNally and Vale, 1993). In this scenario, short microtubule seeds are generated which may be reoriented by diffusion forces in either direction (plus-end-in or plus-end-out) with equal probability. Indeed, active bidirectional transport of short microtubule fragments in the axons has been described in cultured rat hippocampal neurons (Liu et al., 2010). The authors proposed that the transport of these short microtubule fragments is driven by dynein. Our prediction is that microtubule fragments transported in the retrograde direction should be oriented with plus-ends toward the cell body while anterogradely transported fragments should have the opposite orientation.

It has been shown in many different model systems that kinesin-1 and dynein are the two major motors involved in reorganizing cytoplasmic microtubules (Fink and Steinberg, 2006; Straube et al., 2006; Jolly et al., 2010; Mazel et al., 2014). Both motors play an important role in axon formation. However, the contribution of each motor likely differs at different developing stages (see model described in Figure 4). In spherically shaped undifferentiated cells, kinesin-1 motors slide antiparallel microtubules perpendicularly to the plasma membrane with their minus-end-leading (Figure 4A). This sliding generates the force that breaks the symmetry of the cell and induces a deformation of the plasma membrane to initiate neurite outgrowth. During this stage, the role of cortical dynein is probably limited by steric restrictions. Cortical dynein can only slide microtubules tangential to the plasma membrane, and therefore dynein-mediated movement cannot deform the membrane and initiate outgrowth. At the next stage, nascent neurites contain antiparallel microtubule arrays (Figure 4B). This configuration allows kinesin-1 to continue to drive antiparallel microtubule sliding. Once the processes start to form, the new geometry will allow cortical dynein to contribute to both microtubule organization and neurite outgrowth. Cortical dynein in the neurite cortex can only slide microtubules parallel to the axis of the process. This contribution of cortical dynein likely increases as the processes grow longer (engaging more cortical dynein molecules) and thinner (allowing dynein to reach a larger fraction of microtubules in the processes).

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There are two possible microtubule orientations in the neurite that can interact with cytoplasmic dynein. Microtubules with minus-end-out (microtubule of the ‘wrong’ polarity) will be moved by dynein toward the cell body, therefore eliminating them from the processes. At the same time, dynein can interact with and push plus-end-out microtubules moving them toward axon tips, and thus contributing to the forces that drive axonal growth (Figure 4B). As a result of this continuous sorting activity of dynein, developed axons contain uniform microtubule arrays with their plus-end distal (Figure 4C). Under this new microtubule configuration, kinesin-1 contributes little to microtubule sliding (as antiparallel microtubule arrays are needed for kinesin to engage in sliding), but instead favors its bundling activity (Figure 1A, right panel). In addition to its contribution to neurite outgrowth, cortical dynein may have an important role in axon maintenance, removing nascent microtubules of wrong orientation that may appear in the axon either due to new microtubule polymerization or due to the severing of pre-existing microtubules. Because axons and dendrites are common morphologic features observed in neurons from ancestral metazoans to mammals, we postulate that the same molecular mechanism presented here for axon formation in Drosophila neurons is conserved in other organisms. Of course, this ‘microtubule-centric’ model does not describe all the mechanisms involved in axon formation, and clearly leaves out the critical question of axon guidance, but it provides clear roles for multiple microtubule motors involved in axon formation and microtubule organization.

Microtubule orientation differs between axons and dendrites. While axons are filled with uniformly oriented microtubules with plus-end-out, dendrites contain either microtubules of mixed orientation (mammalian neurons) (Baas et al., 1988) or a majority of microtubules minus-end-out (Drosophila and C. elegans neurons) (Stone et al., 2008; Yan et al., 2013). We speculate that neurons selectively employ cortical dynein to dictate which of the nascent neurites will become the future axon. It is likely that the microtubule sliding activity or efficiency of dynein recruitment may be downregulated in dendrites. This hypothesis is in agreement with a recent study in Drosophila da sensory neurons showing that disruption of NudE, a dynein cofactor, impairs microtubule orientation in axons, without affecting the orientation of dendritic microtubules (Arthur et al., 2015). Furthermore, Yan et al. directly demonstrated that kinesin-1 (unc-116) is required for minus-end-out orientation of microtubules in dendrites (Yan et al., 2013). Those observations support the idea that the activity of cortical dynein is downregulated in dendrites, thus preserving the initial minus-end-out orientation of microtubules created by kinesin. Future experiments are required to unravel how dynein interacts with cortical actin and how dynein’s microtubule-sorting activity is regulated in neurons.

To visualize microtubule minus-ends in the Drosophila S2 cell system, a cDNA encoding mouse CAMSAP3 (Jiang et al., 2014) was cloned into the pMT-GFP and pMT-mCherry backbones by NotI-AgeI restriction enzyme sites to generate GFP-CAMSAP3 and mCherry-CAMSAP3. GFP-CAMSAP3 was also cloned into UASp backbone by KpnI-XbaI to create a transgenic Drosophila fly line containing UASp-GFP-CAMSAP3 by standard P element-mediated transformation. A plasmid encoding EB1-GFP under endogenous EB1 promoter (pMT-EB1:EB1-GFP) was used to visualize microtubule plus-ends in S2 cells. For recruitment experiments, all constructs were cloned into pAC.V2014, a modified version of pAC5.1 containing the following multiple cloning site (KpnI-NheI-BmtI-HindIII-AscI-EcoRI-NotI-XbalI-EcoRV-XhoI). FRB-GFP and PEX3-mRFP-FKBP fragments from pβActin-GFP-FRB and pβactin-PEX3-mRFP-FKBP plasmids (Kapitein et al., 2010) were subcloned in the HindIII-NotI and HindIII-EcoRI sites to generate pAC.V2014-FRB-GFP and pAC.V2014-PEX3-mRFP-FKBP. Drosophila BicD cDNA (LD17129 from DGRC) was amplified by PCR and ligated in the AscI-NotI sites of pAC.V2014-FRB-GFP to create pAC.V2014-FRB-GFP-BicD. FKBP fragment was inserted in pAC.V2014 using the AscI-EcoRI sites to create pAC.2014-FKBP. To recruit the FKBP domain to the membrane, the DNA sequence encoding the transmembrane domain GAP-43 (MLCCMRRTKQVEKNDEDQKI) was ligated in the KpnI-AscI sites of pAC2014-FKBP to create pAC2014-GAP43-FKBP.

Fly stocks and crosses were cultured on standard cornmeal food based on Bloomington Stock Center’s recipe at room temperature. The following fly stock lines were used in this study: UASp-tdEOS2-αtub84B (2nd and 3rd chromosome insertions) (Lu et al., 2013b), ubi-EB1-GFP (3rd chromosome insertion) (Shimada et al., 2006); UASt-EB1-GFP (3rd chromosome insertion) (Rolls et al., 2007); elav-Gal4 (3rd chromosome insertion, Bloomington stock #8760) (Luo et al., 1994); UASp-GFP-CAMSAP3 (2nd chromosome insertion, created in this study); DHC64C-RNAi TRiP lines, (Valium 20, Bloomington stock #36698, 3rd chromosome attP2 insertion, targeting DHC64C CDS 1302–1322; Valium 22, Bloomington stock #36583, 2nd chromosome attP40 insertion, targeting DHC64C CDS 10044–10064). Stocks of yw; DHC64C-TRiP RNAi-Valium22; ubi-EB1-GFP,yw; UASt-EB1-GFP; DHC64C-TRiP RNAi-Valium20, and yw; UASp-GFP-CAMSAP3; DHC64C-TRiP RNAi-Valium20 were generated using standard balancing procedures, and crossed with elav-Gal4 to examine EB1-GFP or GFP-CAMSAP3 in DHC64C knockdown.

Primary neurons were obtained from brains of 3rd instar larva as previously described (Lu et al., 2015). Neurons were plated onto Concanavalin A-coated coverslips in supplemented Schneider’s medium (20% fetal bovine serum, 5 μg/ml insulin, 100 μg/ml penicillin, 100 μg/ml streptomycin, and 10 μg/ml tetracycline). For actin depolymerization assays, neurons were plated in Xpress medium for 2 hr before addition of LatB. Drosophila S2 cells were cultured as previously described (Barlan et al., 2013). To induce the formation of processes in S2 cultures, cells were plated in the presence of 0.5 µM LatB. For knockdown assays in S2 cells, cultures at 1.5 x 10⁶ cells/mL were treated twice with 20 µg of dsRNA (day 1 and day 3) and cell analysis was performed on day 5. Double-stranded RNA was transcribed in vitro with T7 polymerase, and purified using LiCl extraction. Primers used to create T7 templates from fly genomic DNA were as follows. T7 promoter sequences (TAATACGACTCACTATAGGG) were added to the 5′ end of each primer). DHC, forward, AAACTCAACAGAATTAACGCCC; reverse, TTGGTACTTGTCACACCACT (Jolly et al., 2010); p150^Glued, forward GAGTTTGAGGAGACGATGGACCACC, reverse GTTGCACGATGGGGTTTCCTTTGCAG; Lis1, forward GGTTGAATTACGCGATCATGAGCATACTGTGGA, reverse GGAGGTGCAGAAATGCTGATGCGCGTATAG; NudE, forward, GCTCAAGTTGGAATCGCATGGCATCGATATGTC, reverse, CTCTCGTCTCATCCATTAATCGCTGTAGTTTTTCCTGC. To induce the formation of short microtubule fragments, Drosophila S2 cells stably expressing GFP-CAMSAP3 were treated with 25 µM Vinblastine for 1 hr. Soluble fraction of tubulin was removed using BRB80 buffer (80 mM PIPES buffer (pH 6.8), 1 mM EGTA, 1 mM DTT, and 1 mM MgCl₂) supplemented with 1% Triton X-100. Extracted cells were fixed and microtubules were immunostained with mouse anti-α-tubulin (DM1α).

To image EB1 comets, CAMSAP3 localization and lysosome transport in Drosophila S2 cells and primary neurons, a Nikon Eclipse U2000 inverted microscope equipped with a Yokogawa CSU10 spinning disk head, Perfect Focus system (Nikon) and a 100 X 1.45 NA lens was used. Images were acquired using Evolve EMCCD (Photometrics) and controlled by Nikon Elements 4.00.07 software. For EB1 and CAMSAP3 time-lapses, images were collected every 2 s for 1 min (EB1 comets) and every 1 min or 5 min for 16 min or 60 min, respectively (CAMSAP3). To image the plasma membrane, CellMask Deep Red dye (1:10,000) was added in both cultured neurons and S2 cells 5 min before imaging. For To visualize sliding, a small fraction of microtubules in cultured neurons expressing tdEOS-αTub84B was photoconverted. Photoconversion was performed by confining the illumination of a heliophore laser (405 nm) in the epifluorescence pathway using a diaphragm. Images were collected once per 30 s for 5 min. To image the recruitment of BicD to the plasma membrane and the distribution of CAMSAP3 in mature neurons, TIRF images were collected using a Nikon Eclipse U2000 inverted microscope equipped with a Plan-Apo TIRF 100×/1.45 NA objective and a Hamamatsu CMOS Orca Flash 4.0 camera (Hamamatsu Photonics, Hamamatsu, Japan), controlled by MetaMorph 7.7.7.0 software (Molecular Devices, Downingtown, PA). Statistical significance for CAMSAP3 populations was determined using two-tailed Fischer’s test with a confidence interval of 95%.

The orientations of EB1 comets both in cultured neurons and S2 cells were quantified using MATLAB and ImageJ. EB1 comets were tracked using the plus-end tracking algorithm in u-track 2.0, developed by Gaudenz Danuser’s group (Jaqaman et al., 2008; Matov et al., 2010). The x-y positions of EB1 comets were extracted and loaded into a custom ImageJ plugin. This semi-automated plugin defined the center of cells and determined the angle of each comet’s trajectory as compared with the cell center. For curved and serpentine axons, trajectories were subdivided into linear segments before ImageJ analysis to ensure that comet orientation was correctly identified. Comet trajectories with an angle >290° or <70° compared with the cell center were defined as plus-end-out. Comet trajectories with an angle >110 and <250 degrees were defined as plus-end-in. Only comets contained in processes were included in the analysis. To create kymographs presented in Figure 1E and Figure 2A the Reslice plugin developed in FIJI was used. Statistical significance for EB1 comets was determined using the non-parametric Mann-Whitney test with a confidence interval of 95%. This test analysis compares the distributions of two unmatched groups.

For Western blot analysis of S2 cell extracts, the following primary antibodies were used: anti-DHC monoclonal antibody 2C11-2 (Sharp et al., 2000) and rabbit polyclonal antibody against KHC head domain provided by A. Minin (Institute of Protein Research, Russian Academy of Sciences, Moscow, Russia).

The formation of the FRB-FKBP complex in all recruitment experiments was induced by addition of 1 µM rapalog (final concentration) (A/C Heterodimerizer, Clontech) to the culture medium. To test the efficiency of recruitment experiments, Drosophila S2 cells were transiently cotransfected either with plasmids encoding FRB-GFP and PEX3-mRFP-FKBP or FRB-GFP-BicD and GAP43-FKBP. Cells were imaged before and 30 min after addition of rapalog to the medium. To directly recruit endogenous dynein to the membrane, S2 cells were cotransfected with plasmids codifying GAP43-FKBP, FRB-GFP-BicD and mCherry-CAMSAP3 in the following ratio (3:1:1). Cells were plated with 10 µM LatB for 2 hr to allow the accumulation of microtubule minus-ends at process tips. Distribution of mCherry-CAMSAP3 was tracked by time-lapse imaging before and after addition of the drug (to the final concentration of 1 µM).

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

The authors thank the Bloomington Stock Center (NIH P40OD018537) for fly stocks, and Drosophila Genomics Research Center (NIH 2P40OD010949-10A1), S Rogers (UNC Chapel Hill), A Akhmanova and C Hoogenraad (both from Department of Biology, Utrecht University) for plasmids. Research reported in this publication was supported by the National Institute of General Medical Science of the National Institutes of Health under award number R01GM052111.

© 2015, del Castillo et al.

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