Drosophila Katanin is a microtubule depolymerase that regulates cortical-microtubule plus-end interactions and cell migration

Abstract

Regulation of microtubule dynamics at the cell cortex is important for cell motility, morphogenesis and division. Here we show that the Drosophila Katanin, Dm-Kat60, functions to generate a dynamic cortical-microtubule interface in interphase cells. In S2 cells, Dm-Kat60 concentrates at the interphase cell cortex where it suppresses the polymerization of microtubule plus-ends thereby preventing the formation of aberrantly dense cortical arrays. Dm-Kat60 also localizes to the leading edge migratory D17 cells and negatively regulates multiple parameters of their motility. Finally, in vitro, Dm-Kat60 severs and depolymerizes MTs from their ends. Based on these data, we propose that Dm-Kat60 removes tubulin from microtubule ends or lattice that contact specific cortical sites to preventing stable and/or lateral attachments. The asymmetric distribution of such an activity could help generate regional variations in MT behaviors involved in cell migration.

Introduction

Microtubules (MTs) form complex and dynamic arrays with pivotal roles in the development and function of eukaryotic cells. While MTs are intrinsically dynamic, their cellular behaviors are tightly regulated by a host of additional factors^1,2. Thus, the MT cytoskeleton is responsive to a variety of cues and can locally adapt its dynamic properties accordingly. These regulatory inputs appear to be particularly relevant at the cell cortex where localized alterations in MT dynamics and organization are central to cell migration, polarization, morphogenesis, and division^3–6.

Katanin is a phylogenetically conserved enzyme which utilizes the energy of ATP hydrolysis to generate MT breakage, in vitro⁷. Katanin was originally purified from sea urchin eggs as a heterodimer of a 60 kD catalytic subunit (p60) and an 80 kD targeting and regulatory subunit (p80)⁸. Katanin p60 and p80 homologues have now been identified in evolutionary diverse systems and many organisms contain several genes encoding distinct p60 and/or p80 proteins. Functional analyses reveal diverse roles for Katanin in mitosis/meiosis^9–12, neuronal morphogenesis^13,14, and the assembly/disassembly of cilia/flagella^15–18. In addition, a Katanin in higher plants has been shown to regulate the assembly of cortical MT arrays which, in turn, determine the directional deposition of cellulose and thus impact cell morphogenesis^19–22. In this context, Katanin releases new MTs nucleated from the walls of pre-existing MTs^7,23.

We previously found that the Drosophila Katanin p60, Dm-Kat60, associates with mitotic chromosomes and stimulates the depolymerization of kinetochore-associated MT plus-ends during anaphase A¹². In the present study, we tested the hypothesis that Dm-Kat60 also functions to regulate MT dynamics during interphase - a topic that has not been addressed in any other animal system.

Results

Dm-Kat60 localizes to the interphase cell cortex

We first probed Dm-Kat60’s localization in cultured interphase S2 cells using immunofluorescence. Dm-Kat60 immunostaining was found to be strongly enriched within a 0.5–3 μm thick band extending around the entire cell boundary (Fig. 1a). This cortical staining pattern was reduced or abolished by Dm-Kat60 RNAi supporting its specificity (see Supplementary information, Fig. S1a). Ectopically-expressed Dm-Kat60-GFP also acquired a cortical localization (see Supplmentary information, Fig. S2 a,b).

Figure 1. Dm-Kat60 targets to the interphase cell cortex.

(a) Immunofluorescence micrograph showing the localization of Dm-Kat60 in interphase S2 cells (antibody characterized in¹²). (b) Immunofluorescence micrograph of an interphase S2 cell double-labeled for MTs (anti-α-tubulin; red) and Dm-Kat60 (green). (c) High magnification of the region boxed in b. (d) Immunofluorescence of an interphase S2 cell double labeled for actin (red) and Dm-Kat60 (green). Higher magnification of the two regions boxed in the “merge” panel. Scale bar, 10 μm (panels a,b,d) or 2 μm (panel c).

Though Dm-Kat60 showed little co-localization with MTs in the interphase cell interior, numerous MT plus-ends were observed to extend into and terminate within the Dm-Kat60-rich cortical zone (Fig. 1b,c). Dm-Kat60 also generally co-localized with cortical actin (Fig. 1d). Dm-Kat60’s cortical localization occurs independently of MTs as it persists following MT depolymerization with colchicine. However, it is nearly completely abolished when cells are treated with cytochalasin D indicating a strong reliance on actin (see Supplementary information, Fig. S3). On the other hand, Dm-Kat60 RNAi did not prevent the cortical accumulation of actin (see Supplementary information, Fig. S1b).

Dm-Kat60 regulates interphase MTs

We then performed live analyses of MT behaviors in control and Dm-Kat60 RNAi-treated S2 cells expressing GFP-α-tubulin. In controls, MTs generally grew perpendicularly to the cortex, briefly paused upon contacting the cortex, and then catastrophed (transition from growth to shrinkage) into the cell interior (Fig. 2a; Movie S1a; see also ²⁴). >90% of plus-end catastrophes (78/84 observed in 9 cells) occurred within 3 μm of the cell edge indicating the Dm-Kat60-rich cortical zone as a potential “hot-spot” for the induction of MT depolymerization.

Figure 2. Depletion of Dm-Kat60 causes significant MT curling and bundling beneath the cortex.

(a) Confocal images of live GFP-tubulin expressing S2 cells treated with control RNA. The cell border is marked with a dashed red line in the higher magnification image shown in the bottom right. The bottom right panel is also shown in Supplemental movie s1a. (b) Time series of a putative MT severing event at the cortex. The green arrow marks the site of MT breakage and the dashed red line marks the cell border. See also, Supplemental movie S1c. (c) Images of live GFP-tubulin expressing S2 cells treated with Dm-Kat60 RNAi. As in (a), the cell border is marked with a dashed red line in the higher magnification image shown in the bottom right of this panel. This panel is also shown in Supplemental movie S1b. (d) Time series of images obtained from a Dm-Kat60 RNAi treated cell showing the lateral growth of MT ends along the cortex and their subsequent incorporation into the underlying MT array. Individual ends are indicated by colored circles and time (seconds) is labeled in each image. Scale bar, 10 μm (panels a, c), 1 μm (panel b) or 2 μm (panel d).

Interestingly, some MTs were also observed to break very near their ends upon contacting the cortex. The newly formed plus-ends of these “severed” MTs initiated catastrophe immediately thereafter (Fig. 2b; Movie S1c). While such events were rarely observed when cells were imaged at 5 second intervals (our standard for longer time-series), they appeared much more frequently when images were acquired at subsecond intervals (0.2–0.8 images/second). Under these conditions, putative severing events near MT tips preceded ~30% of observed catastrophes (20/63 microtubule catastrophes observed in 7 cells).

In stark contrast to controls, MTs in the majority of Dm-Kat60-depleted cells were noticeably curled beneath the cortex and often formed dense parallel arrays ringing the cell periphery (Fig. 2c; Movie S1b). This phenotype was largely reversed by the induced expression of an RNAi-resistant Dm-Kat60-GFP construct (see Supplementary information, Fig. S2c,d). While we were unable to observe the formation of these cortical arrays de novo, examination of the few clearly visible individual MTs present at the onset of imaging provided insights into their genesis. Similarly to controls, these MTs often contacted the cortex plus-end on. However, instead of rapidly initiating catastrophe, they continued to grow along the cortex until coming into proximity of the underlying ring of MTs, into which they were ultimately incorporated (Fig. 2d).

A unique role for Dm-Kat60 in MT dynamic instability

Roughly a quarter of the Dm-Kat60 RNAi-treated interphase S2 cells imaged contained at least some cortical domains with numerous unambiguously identifiable MT plus-ends. The behaviors of these were analyzed in detail. To measure large numbers of plus-end behaviors, we developed an automated tracking algorithm which could accurately follow the position of MT ends over time (Fig. 3a; Movie S1d,e; see Methods). The validity of this approach was confirmed by comparing a subset of MT trajectories generated automatically to those generated by hand (see Supplementary information, Fig. S3).

Figure 3. Automated tracking and quantitative analysis of MT plus-end organization and dynamics in control and Dm-Kat60 RNAi treated cells.

(a) Ends of GFP-labeled MTs were tracked with an automated tracking algorigthm. Each tracked MT end is marked with a colored dot: growing ends in blue, pausing end in yellow, and shrinking ends in pink. The stack of panels on the right is a time series of the boxed region.

(b) Trajectories of MT ends tracked in a control RNA-treated cells cell. Individual MT trajectory paths are indicated by colored lines. (c) Trajectories of MT ends tracked in a Dm-Kat60 RNAi cell. Again, individual trajectory paths are indicated by colored lines. Relative to controls, significantly more MTs continue to grow after reaching the cortex, and so MTs are frequently observed to bend and grow parallel to the cortex. (d) Compared to control cells (7 cells, 257 MT, 2600 distances), the MT ends in Dm-Kat60 RNAi cells (5 cells, 494 MTs, 6087 distances) spent significantly (p < 0.0001) more time in the vicinity of the cortex. MT ends within the cell interior could not be identified and thus were not analyzed. (e) Cortical MTs spend more time growing after Dm-Kat60 knockdown. The change in dynamic behavior is most prominent in MT ends very near (< 1.3 μm) the cell margin, but is still significant (p < 0.0001) for ends further (> 1.3 μm) away. Numbers within bars indicate N for cells, and N for MT ends. Error bars, SEM. Scale bars, 10 μm (all image panels).

Consistent with our visual inspection of the data, in controls, the majority of MT plus-end growth/shrinkage trajectories occurred along largely straight paths oriented perpendicular to the cortex (Fig. 3b). By contrast, Dm-Kat60 RNAi cells displayed numerous plus-end trajectory paths parallel to the cortex (Fig 3c). Moreover, on average, MT plus-ends in Dm-Kat60 RNAi cells spent significantly more time in growth (as opposed to shrinkage) parallel to the cortex and were positioned significantly closer to the cell edge (Fig. 3d,e).

We then analyzed the life history plots of the tracked plus-ends to specify Dm-Kat60’s impact on dynamic instability (Table 1). As expected, Dm-Kat60 RNAi significantly suppressed plus-end catastrophes >2-fold. Thus, Dm-Kat60 is normally a potent catastrophe promoter. Dm-Kat60 depletion also increased transitions from pause to growth; paused plus-ends were almost twice as likely to rescue to the growth state. Surprisingly, the frequency of the reverse transition (a growing plus-end stalling into pause) also increased after Dm-Kat60 RNAi, but by a more modest 20%. Finally, we measured a slight (~20%) but significant increase in MT growth rate in Dm-Kat60 RNAi cells. On the other hand, depletion of Dm-Kat60 did not significantly impact the frequency of transitions out of the shortening state nor did it alter the average rate of shrinkage.

Table 1.

Comparison of state-to-state frequency (#/sec) and growth/shrinkage rate (nm/sec) between control, Dm-Kat60RNAi and Klp10A RNAi cells

State-to-state	Control RNA	Dm-Kat60 RNAi	Klp10A RNAi
G → S	0.0106 (34) a	0.0041 (7)	0.0028 (4)
	0.0072–0.0144 b	0.0017–0.0081	0.0007–0.0073
		p=0.0249 *c	p=0.0004 ***
G → P	0.0479(167)	0.0576 (75)*	0.0958 (116)
	0.0432–0.0522	0.0501–0.0645	0.0809–0.1053
		p=0.0225 *	p=0.0181 *
S → G	0.0098 (21)	0.0110 (10)	0.0045 (2)
	0.0063–0.0146	0.0056–0.0195	N/A
		p=0.7687	N/A
S → P	0.0521 (109)	0.0590 (42)	0.1113 (47)
	0.0466–0.0575	0.0477–0.0678	0.0972–0.1242
		p=0.2084	p=0.0035 **
P → G	0.0291 (72)	0.0481 (47)	0.0641 (60)
	0.0223–0.0369	0.0360–0.0627	0.0461–0.0817
		p=0.0145 *	p=0.1778
P → S	0.0216 (77)	0.0156 (22)	0.0119 (14)
	0.0166–0.0275	0.0080–0.0263	0.0059–0.0237
		p=0.2657	p=0.0031 **
Growth	52.79 (1083)	62.13 (580)	58.68 (530)
	1.25 d	1.97	1.73
		p<0.0001 ****	p=0.033 *
Shrinkage	84.88 (819)	81.96 (316)	58.43 (179)
	2.81	4.61	3.63
		p=0.8380	p=0.0001 ***

^aNumbers in parentheses indicate the event count in control RNA (5 cells), Dm-Kat60 RNAi (7 cells) or Klp10A RNAi (5 cells) treatment.

^bIntervals indicated are 95% confidence intervals calculated using bootstrapping procedure with 1000 resamples, and BCa correction.

^{c *}indicates statistically different from control if p < 0.05 (p values were shown).

^**if p < 0.01,

^***if p < 0.001, and

^****if p < 0.0001. Statistical significance determined using a simulated two-tail permutation test with 1000 resample.

^dSEM.

The Drosophila Kinesin-13, KLP10A, has also been found to promote catastrophes at cortex of interphase S2 cells²⁴. To better understand Dm-Kat60’s functional inter-relationship with this protein, we reassessed KLP10A’s influence on MT dynamics using our current methodologies (Table 1). As with Dm-Kat60 RNAi, the depletion of KLP10Asignificantly reduced catastrophe frequency. However, KLP10A RNAi also increased the frequency of transitions into pause--Kat60 RNAi had the opposite effect. Also unlike Dm-Kat60 RNAi, KLP10A RNAi had no influence on MT growth rate but strongly suppressed the rate of shrinkage.

Dm-Kat60 localizes to the leading edge of D17 cells and regulates their migration

Dm-Kat60’s influence on MT dynamics at the cell cortex suggested a role in cell migration^25–28. Since S2 cells are immotile, we examined this possibility in migratory D17 cells. D17 cells were originally isolated from cultures of dissected imaginal discs²⁹; despite their epithelial origin, D17 cells exhibit a gene expression profile that is consistent with that of Drosophila hemocytes (data not shown). Like hemocytes^30–32, D17 cells spontaneously polarize in culture, assemble an exaggerated leading edge, and exhibit robust cell motility (Movie S2). Immunofluorescence revealed that Dm-Kat60 accumulates at the leading edge of polarized D17 cells where it extensively co-localizes with actin (Fig. 4a). The other MT-severing enzymes in Drosophila, Spastin and Fidgetin^12,33, displayed no such cortical enrichment nor did their depletion obviously alter D17 cell migration (see Supplementary information, Fig. S5).

Figure 4. Dm-Kat60 targets to the leading edge of motile D17 cells and negatively regulates their migration.

(a) Immunolocalization of Dm-Kat60, tubulin, and actin in polarized Drosophila D17 cells. The arrowheads indicate enrichment of Dm-Kat60 at the leading edge; the boxed areas are magnified in far right panels. (b) Western blot showing the depletion of Dm-Kat60 in D17 cells using RNAi (see also Supplemental information, Fig. S8b). (c) Representative phase-contrast images of scratch-wounds at 0 hr and 24 hr after wounding.. (d) Quantification of migration during wound closure for control and Dm-Kat60 RNAi-treated D17 cells. Migration area was calculated by subtracting the total wound area at 24 hr from the total wound area at 0 hr after wounding and then normalized to control RNA. p = 8.5 × 10⁻⁵. Data represent mean values ± SEM from four independent experiments. Numbers in bars are sample sizes. Scale bars, 10 μm (panel a) or 50 μm (panel c).

Dm-Kat60’s influence on D17 cell migration was first examined in a wound healing assay. D17 cell cultures were treated with Dm-Kat60 or control dsRNA for 10 days, the confluent monolayers were wounded with a pipette tip, and the resulting “wounds” were imaged immediately and 24 h later. Unexpectedly, we measured a significant 1.4-fold increase in the rate of wound closure following Dm-Kat60 RNAi (Fig. 4b–d). To specify the basis of this difference, we then assessed the intrinsic motility properties of single cells using time-lapse microscopy (Fig. 5a, Movie S2). Dm-Kat60 RNAi-treated cells moved significantly faster than controls with a striking increase in the number of high velocity movements (Fig. 5b,c). They also exhibited an increase in total migration distance (Fig. 5d). Although control D17 cells do not exhibit directional motility in this assay, we did observe a slight, but significant decrease in directionally persistent migration (Fig. 5e), suggesting that the increased wound migration of Dm-Kat60 depleted cells does not result from increased directionality. Finally, we measured a notable difference in persistent migration following Dm-Kat60 RNAi. Whereas migrating control cells spent ~63% of their time moving, cells depleted of Dm-Kat60 spent ~73% of their time migrating (Fig. 5f). Taken together, these data suggest that Dm-Kat60 normally serves as a negative regulator of cell motility by suppressing fast and persistent migration.

Figure 5. Dm-Kat60 negatively regulates multiple parameters of D17 cell motility.

(a) Representative migration tracks of 20 cells for each treatment group (control and Dm-Kat60 RNAi; also see movie S2). (b) Frequency distributions of instantaneous cell velocities from time-lapse imaging of individual D17 cell movements. p = 8.2 × 10⁻⁴⁰.

(c) Quantification of the ratio of velocity frequencies between Dm-Kat60 and control RNAi treatments. The Dm-Kat60/control RNAi ratio was calculated by dividing the number of movements for Dm-Kat60 RNAi-treated cells by the number of movements for control RNA-treated cells for a given range of velocities from the graphs in b. A ratio less than one (red) represents a decreased number of movements for Dm-Kat60 RNAi relative to control RNA, and a ratio greater than one (blue) represents an increased number of movements for Dm-Kat60 RNAi relative to control RNA. Relative to control, Dm-Kat60 depletion decreases the frequency of cells migrating at slow rates while increasing the frequency of fast migration rates.

(d) Quantification of total migration distance over a 3 hr time period. p = 1.5 × 10⁻⁵. (e) Quantification of intrinsic cell directionality. Directionally persistent migration (D/T) was calculated as a ratio of direct distance between start and end points (D) to the total migration distance (T). p = 3.6 × 10⁻².

(f) Quantification of persistent migration. The percentage of time cells spent migrating was calculated by subtracting those movements between frames that were less than 2 μm (considered to be migratory pauses) from the total number of movements and then dividing by the total number of movements. p = 3.7 × 10⁻⁵ Data represent mean values ± SEM from four separate dsRNA treatments. Numbers in bars are sample sizes.

Does Katanin regulate the migration of human cells? While conventional human p60 had no apparent impact on cell movement (not shown), the p60-like protein, KATNA1 (KL1)^34,35, does localize to the leading edge of migratory cells, regulate MT growth and negatively impact cell movement (see Supplementary information, Fig. S6a–e). Thus, human KATNA1 is likely the functional orthologue of Dm-Kat60, at least during interphase.

Dm-Kat60 inhibits actin protrusions at the cell edge

MTs are believed to impact cell movement largely through the regulation of actin based structures, such as lamellae^5,36–39. Unfortunately, technical limitations precluded us from testing these cytoskeletal interactions in D17 cells. However, this could be studied in S2 cells co-expressing GFP- α-tubulin and mCherry-actin. While Dm-Kat60 RNAi did not significantly impact on the accumulation of actin at the cortex or the rate of actin retrograde flow (not whosn), it did strongly influence the cycles of lamellipodium protrusion and retraction. Specifically, Dm-Kat60 RNAi treated cells displayed a significant ~3-fold increase in the frequency of protrusions and >2-fold increase in the average displacement per protrusion (Fig. 6b,c; Movie S3). A localized increase in the protrusion/retraction cycle at the leading edge could conceivably underlie the increased cell migration rates observed following Dm-Kat60 RNAi.

Figure 6. Dm-Kat60 negatively regulates actin protrusions at the cell edge.

(a) Time lapse images showing the cortical dynamics of control and Dm-Kat60 RNAi-treated S2 cells expressing GFP-α-tubulin and mcherry-actin. Yellow lines outline the edges of the cortical actin arrays in each image. These lines are stacked vertically (and shown in red) at the far right to illustrate the time-dependent alterations in the morphology of the cell edge (protrusions) in each condition (T shows time axis). (b) Representative kymographs of mcherry-actin labeled cortical regions from a control and Dm-Kat60 RNAi-treated cell. (c). Dm-Kat60 RNAi significantly increases both the frequency (left) and average displacement (right) of actin-based membrane protrusions ( p<0.0001 for both). Data represent Mean ± SEM and numbers in the columns indicate regions/cells analysed. Scale bars, 2 μm for both panel a and b.

Kat-60 is MT severing enzyme and end depolymerase

Finally, we examined Dm-Kat60’s in vitro mechanism-of-action using purified, baculovirus-expressed, GFP-Dm-Kat60 (rDm-Kat60). Taxol-stabilized, rhodamine-labeled MTs were immobilized on coverslips⁴⁰, incubated with 50 nM rDm-Kat60 (Fig. 7a) and imaged through time using total internal reflection microscopy (TIRF). rDm-Kat60 robustly severed taxol-stabilized MTs in the presence of ATP (Fig. 7b, c; Movie S4) and this reaction was inhibited when ATP was depleted with hexokinase and glucose or replaced with the non-hydrolyzable ATP analogue, AMPPNP (Fig. 7e). Thus, Dm-Kat60-mediated severing is stimulated by ATP hydrolysis. Severing also likely involves an interaction between Dm-Kat60 and the C-terminal tail of tubulin since it was almost completely inhibited by the protease subtilisin, which removes the C-termini of the tubulin subunits within the MT polymer (not shown). Subtilisin treatment was previously reported to inhibit Katanin-mediated severing of MTs⁸.

Figure 7. Dm-Kat60 severs and depolymerizes MTs from their ends.

(a) Coomassie-stained SDS-PAGE gel of the purified rDm-Kat60 protein used for analysis (see also Supplemental information, Fig. s8c). Arrow indicates the band corresponding to rKat60.

(b) Time-series of TIRF images showing the disassembly of a field of immobilized MTs by 50 nM rDm-Kat60 and ATP. Green arrows mark the ends of an individual MT at the beginning of visualization; a red arrow marks a MT end that has shortened from its initial position. Time (seconds) is indicated.

(c) (Left) Time-series of an individual MT (marked by arrows in c) before (pre) and after addition of rDm-Kat60. i–iii kymographs showing the depolymerization and severing (indicated by dotted yellow lines) of individual MTs from other experiments. The time (t) axis is vertical and distance (xy) axis horizontal in all kymographs shown in this figure

(d) (Left) Time-series of an individual polarity marked MT incubated with rDm-Kat60. The plus-end is dimly labeled and the minus-end is brightly labeled. i & ii. Kymographs showing the depolymerization and severing (dotted yellow lines) of additional polarity marked MTs.

(e) Upper: Quantification of the frequency of MT severing by rDm-Kat60. All conditions included 2 mM ATP (or 2 mM AMPPNP). P = 0.031

Lower: Measured rates of MT plus- and minus-end depolymerization by rDm-Kat60. p = 3.1 × 10⁻⁷. Again, all conditions included 2 mM ATP or AMPPNP. Data in both panels represent mean ± SEM. N= # of MTs analyzed.

(f) Electon micrographs showing the ends of negatively stained MTs after incubation with: no Dm-Kat60 (control), 50 nM rDm-Kat60 and 2mM ATP (Dm-Kat60), or full-length KLP59D and ATP (KLP59D).

(g). Quantification of MT ends per unit MT length. Compared with the no Dm-Kat60 and Dm-Kat60 (50 nM) + AMPPNP (2mM) controls, Dm-Kat60 + ATP (50 nM GFP-Dm-Kat60, 2 mM ATP) induces 3-fold more ends (p < 0.0001). Numbers in the column indicate numbers of MTs/EM fields. Data represent mean ± SEM pooled from two independent experiments.

Scale bars, 500 μm (panel a), 10 μm (panel b) or 20 nm (panel f).

To our surprise, we also observed substantial rDm-Kat60-induced depolymerization of MT ends which generally preceded any obvious severing of the lattice (Fig. 7b, c). Like severing, end-depolymerization was suppressed by hexokinase and glucose, AMPPNP and subtilisin suggesting that both occur by a similar mechanism (Fig. 7e). While both MT ends were depolymerized by rDm-Kat60, one end almost always depolymerized faster than the other (Fig. 7c). When these assays were performed using polarity marked MTs, the plus-end was found to depolymerize ~3-fold faster than the minus-ends (Fig. 7d, e). To ensure that MT end depolymerization did not result from a protein contaminant, identical analyses were performed using the catalytically inactive Dm-Kat60 (E393Q)⁴¹ mutant. This mutant was entirely incapable of severing MTs and, while it did induce some end depolymerization, the rate at which this occurred was ~5-fold slower than that induced by the wild type protein (Fig. 7e).

In a last set of experiments we examined the morphology of MT ends severed/depolymerized by rDm-Kat60 using electron microscopy (EM). Samples of MTs incubated with rDm-Kat60 and ATP, rDm-Kat60 and AMPPNP, and a no rDm-Kat60 control were applied to EM grids and fixed by negative staining (Fig. 7f). We measured a ~3-fold increase in the number of MT ends/unit length in the rDm-Kat60/ATP condition relative to the AMPPNP and no-rDm-Kat60 controls (Fig. 7g). Thus, rDm-Kat60 was active in these assays. Interestingly, we observed no substantial difference in the morphology of MT ends among any of the above conditions. In all cases, MT ends were blunt or had a few straight protofilaments protruding from the end. By contrast, MTs incubated with the Drosophila kinesin-13, Klp59D⁴², displayed numerous curled protofilaments peeling away from the ends (Fig. 7g) which is typical of kinesin-13 catalysed MT depolymerization⁴³. This suggests that that Dm-Kat60 and kinesin-13s remove tubulin from MT ends via distinct mechanisms

Discussion

Our results identify Dm-Kat60 as a novel and important regulator of microtubule dynamics and cell migration. The human Katanin, KATNA1, behaves similarly. In addition to its cellular roles, in vitro analyses indicate that Dm-Kat60 has the capacity to function both as a MT severing enzyme and end-depolymerase which, to our knowledge, is an unprecedented observation. Based on its cortical localization, RNAi phenotypes, and catalytic activity, we propose that Dm-Kat60 (and KATNA1) contributes to the generation of a dynamic interface between the MT cytoskeleton and the interphase cortex by removing tubulin subunits from any region of the MT making contact Dm-Kat60-rich cortical sites (see Supplementary information, Fig. S7).

Among the more unexpected outcomes of this study is the observation that Dm-Kat60 induces MT end depolymerization, in vitro. However, given current models of Katanin’s interaction with the MT, such a finding is not entirely surprising. Biophysical and biochemical studies have suggested that severing by Katanin is mediated by the transient hexamerization of p60 proteins about the C-terminus of a single tubulin within the MT^7,41. ATP-hydrolysis and/or the subsequent disassembly of the hexamer is believed to generate a mechanical force which, through multiple iterations, induces the removal of the tubulin from the lattice. If Katanin works by “tugging” on a single tubulin heterodimer, then the exposed tubulins at the microtubule end are likely to be the easiest to remove because they lack a longitudinal contact. However, we cannot rule out the possibility that Dm-Kat60-mediated end-depolymerization is a manifestation of multiple severing events occurring very near the tip.

Within the cell, Dm-Kat60’s severing and depolymerase activities likely remain under very tight spatial constraints. In this regard, the recruitment of Dm-Kat60 to the cell cortex appears to be central to its interphase functions. While our data suggest that this process is reliant on the presence of actin but not MTs, the specific mechanisms that deliver Dm-Kat60 to the cortex remain a mystery. One appealing hypothesis is that Dm-Kat60 is directly or indirectly linked to the cortical actin array via Drosophila p80. The p80 subunit contains repeated WD40 motifs known to mediate protein-protein interactions⁴⁴; similar motifs have been identified in some actin-binding proteins⁴⁵. p80’s WD40 repeats are essential for the centrosomal targeting of Katanin in other organisms^46,47. It has also been suggested that p60 acts independently of p80 in some circumstances⁴⁸. The identification of Dm-Kat60’s binding partners represent an important next step in understanding its cellular activities.

At the cortex, Dm-Kat60 suppresses MT growth primarily by inducing plus-end catastrophes and transitions from growth to pause. While other classes of proteins are known to induce MT depolymerization, in vitro^43,49–51, Dm-Kat60’s presumptive ability to remove tubulins from any region of the MT—end or lattice—may be particularly useful in the more complex cellular environment. For example, such an activity could allow Dm-Kat60 to prevent sustained MT growth along the cortex regardless of whether the MT contacts the cortex end-on or side-on. The newly created plus-end at the cortical MT interface would then initiate catastrophe or enter the pause state depending on its association with other MT binding proteins (see below). Moreover, the ends of polymerizing MTs in cells are often “capped” by plus-end binding proteins such as EB1⁵². Dm-Kat60 could remove these by severing the MT at the base of the EB1 “cap” and/or directly removing EB1-bound tubulins from the plus-end. The acidic tail of EB1 could mimic the C-terminus of tubulin thereby providing a substrate for Dm-Kat60⁵³.

Of course, Dm-Kat60 is not alone in its ability to stimulate the catastrophe of MT plus-ends near the cortex of interphase S2 cells as the Drosophila kinesin-13, KLP10A also displays this activity²⁴. However, aside from the ability of both proteins to promote catastrophes, the activities of Dm-Kat60 and KLP10A are quite distinct. A most notable difference is that KLP10A does not concentrate on the cortex but instead binds to the ends of polymerizing MTs to which it is recruited by EB1. Intriguingly, recent work suggests that EB1 can inhibit the depolymerase activity of Kinesin-13s by shielding the plus-end⁵⁴. If Dm-Kat60 were to generate plus-ends lacking EB1, then it could relieve this inhibition. .

We propose that Dm-Kat60 and KLP10A work together as follows: 1) Dm-Kat60 removes tubulins (EB1-bound or otherwise) from regions of the MT that come in close proximity to the cortex thereby creating a free plus-end at the cortical interface. Many of these newly created plus-ends immediately enter a paused state (depletion of EB1 has been shown to strongly promote pause⁵⁵). 2) Next, KLP10A, which has already been accumulated near the end by EB1, promotes the transition of this end from pause to shrinkage—this transition can occur rapidly and may often appear as a catastrophe. Our current study also suggests that KLP10A increases the rate of plus-end depolymerization and thus a small, difficult to detect, portion of the protein may remain associated with the end as it depolymerizes. Why such an effect was not noted in our initial analysis of KLP10A is unknown but may be due to the more limited region of the cortex analyzed in that study²⁴.

The finding that Dm-Kat60 targets to the leading edge of motile D17 cells and alters their migration provides a broader biological context through which our findings can be viewed and interpreted. While Dm-Kat60’s depletion had no obvious influence on the establishment cell polarity, it did increase both the frequency and displacement of membrane protrusions, at least in S2 cells. The localized suppression of protrusions at the leading edge of motile D17 cells could exert negative control over the rate and persistence of cell movement.

The observation that Dm-Kat60 RNAi results in faster and more persistent migration seems consistent with other studies demonstrating that growing MTs stimulate the GTPase, Rac, at the leading edge, which may promote adhesion complex remodeling needed to drive and sustain protrusions^38,39. In addition, because Dm-Kat60 depletion at the leading edge of migratory cells should decrease catastrophes, MTs could become unusually persistent and abundant in the extending protrusion, which may intensify other processes that favor protrusion, like increased kinesin-mediated delivery of vesicles to the protrusion zone⁵⁶.

How does this relate to the organism? A recent study examining hemocyte migration in developing Drosophila embryos, for which we believe D17 cells to be a model, showed that hemocytes migrate less efficiently in response to guidance cues following the disruption of MT dynamics³⁰. Under these conditions, the authors also observed an increase in cell velocities and decrease in directional persistence - similar to D17 cells depleted of Dm-Kat60 by RNAi. Thus, Dm-Kat60 at the leading edge may modulate MT dynamics to “fine-tune” cell migration by suppressing protrusions.

In conclusion, the findings of this study uncover new and unexpected roles for the Drosophila Katanin p60, Dm-Kat60, in the regulation of cortical MT dynamics and provide insights into how the MT cytoskeleton feeds into cell migration. The ability of cells to move and change shape is central to organismal development and when defective have been linked to human diseases such as cancer. Given this, our finding that human KATNA1 performs many of the same functions as Dm-Kat60 suggest the former as a potentially useful new therapeutic target.

METHODS

S2 cell culture, RNAi and construction of plasmids

S2 cells expressing GFP-α-tubulin were a gift from R. Vale (University of California, San Francisco, CA). Wild-type S2 cells were obtained from ATCC (Manassas, VA). Our methods for S2 cell culture, RNAi treatment of S2 cells, and production of dsRNA from the 3′ UTR of endogenous Dm-Kat60 (including sequences) were described previously¹².

Immunofluorescence microscopy

S2 cells were plated on concanavalin A-coated coverslips for 1.5 hrs, fixed in 100% methanol at −20°C for 20 min and blocked with 5% normal goat serum in PBS containing 0.1% Triton X-100. Primary antibodies (Dm-Kat60, α-tubulin [DM1a; Sigma-Aldrich, St. Louis, MO], and actin [clone C4, Millipore, Billerica, MA]) were applied at 1–20 μg ml⁻¹ final concentration in blocking buffer. Fluorescent secondary antibodies (Jackson Immuno-Research Laboratories, West Grove, PA) were used at 7.5 μg ml⁻¹. Imaging was performed on an Ultraview spinning-disk confocal system (PerkinElmer, Waltham, MA) mounted on an inverted microscope (Eclipse TE300; Nikon) with a 100× (1.4 NA) or 60× (1.4 NA) objective and captured with an Orca ER digital camera (Hamamatsu, Japan). For MT disruption, cells were treated with 50 μM colchicine for 18 hrs prior to fixation. For microfilament disruption, cells were treated with 5 μM cytochalasin D for 1 hr before fixation.

Live cell imaging and automated tracking

4D time-lapse movies of GFP-α-tubulin-expressing S2 cells were acquired using the Ultraview spinning disk confocal microscope described above. MT ends were tracked through each series of time-lapse images with an automated tracking program using the following assumptions:

Definition of the tracking region

For each cell, we tracked MTs in a hand-selected region-of-interest (ROI) consisting of an outer (the cortex; Fig. 3a) and an inner (to avoid tracking into the crowded interior) boundary. For all calculations, the data from cells of the same treatment (control or Dm-Kat60 RNAi) were pooled.

Definition of growth and shrinkage

To define growth and shortening, we calculated the angle α between the direction of the displacement of the MT tip and the direction of the MT itself. A value of cos(α)>0.2 (α<~78°) indicated probable growth, and cos(α)<−0.2 (α>~101°) indicated probable shortening. These “loose” cutoffs were chosen to deal with the curvature of MTs in the Dm-Kat60 RNAi cells. To qualify as growing or shortening, we required that the MT display the same behavior (either growing or shortening) for at least two consecutive frames, and that the overall displacement in a run of growth or shrinkage be at least 5 pixels.

Definition of the distance to the cortex

Distance to the cortex was defined as the Euclidean distance from a MT tip to its closest point on the cortex. The difference in the distance distributions was tested using the Kolmogorov-Smirnov two-distribution test.

Definition of growth parallel to the cortex

For two consecutive frames t and t+1, the points on the cortex closest to the trajectory segment are given by c(t+1) and c(t), respectively, and the displacement vector along the cortex is δc(t)=c(t+1)−c(t). A MT was considered to grow parallel to the cortex if the angle between the tip displacement and δc(t) was less than 45°. Error bars plotted are the asymptotic standard errors for a binomial distribution: $\sqrt{p(1-p)/N}$, where p is the fraction that grow along the cortex, and N is the total number of data points. Significance was tested by χ² using a 2×2 table of dichotomous outcomes.

Calculating transition rates

Catastrophes near the cortex were determined by counting the number of significant shortening events away from the cortex. To be counted, a shortening event had to begin within 0.63 μm of the cortex (10 pixels), and increase its distance from the cortex by at least 0.32 μm (5 pixels) by the end of the shortening run. The catastrophe statistic was calculated as

$$\frac{1}{N}\sum _{i=1}^N\left[\frac{[\text{depolymerization}\text{events}\text{in}\text{cell}i]/[\text{s}\text{in}\text{cell}i]}{[\text{microtubules}\text{near}\text{cortex}\text{in}\text{cell}i]}\right]$$

The standard error was estimated by bootstrap sampling on the terms of the sum with 1000 resamples. Statistical significance was determined using 1000 resamples of a one-sided permutation test with the test statistic log(S_1,i/S_2,i), where S_1,i and S_2,i are the catastrophe statistic sums for resample i of control (1) and Dm-Kat60 RNAi (2), respectively. To calculate transition rates (Table 1), each frame of a MT trajectory was marked as either grow, shorten, or pause. Grow and shorten states were defined as previously described. Pause was defined as neither growing nor shortening. The lifetimes of states that began and ended in the middle of a trajectory (not on a boundary), as well as the state they transitioned to, were collected. The rate of transition between an initial state k (k=grow, shrink, or pause) and a final state l (l≠k) is

$$\frac{[\text{no}.\text{of}\text{transitions}\text{from}k\text{to}l]}{[\text{no}.\text{of}\text{transitions}\text{out}\text{of}k]}\times \frac{1}{[\text{no}.\text{of}\text{transitions}\text{out}\text{of}k]}\times \sum _i\left(\frac{1}{[\text{time}\text{in}\text{state}k\text{before}\text{transition}i]}\right)$$

We used a bootstrap approximation to construct 95% confidence intervals of each transition, with 1000 resamples of all lifetimes. Statistical significance was established using a two-sided permutation test with 1000 resamples of the statistic r_1,i – r_2,i, where r_1,i and r_2,i correspond to the rates calculated in resample i of control (1) and Kat60-RNAi (2).

Purification of recombinant 6xHis-GFP-Dm-Kat60

6xHis-GFP-Dm-Kat60 (rDm-Kat60) was cloned into pFastBac HTa (Invitrogen) and then transformed into DH10Bac E. coli cells. The primers to amplify GFP were: forward—GAATTCACCATGGTGAGCAAGGGCGAGGAG and reverse—AAGGAAAAAAGCGGCCGCCTTGTACAGCTCGTCCATGCCGAG. The primers to amplify Dm-Kat-60 were: forward—AAGGAAAAAAGCGGCCGCCACCATGTCCATAACCTTACTGCGAGGTGG and reverse—CCGCTCGAGTCATGACGATCCGAACTCCCTC.

The bacmid was isolated from DH10Bac E. coli and then transfected into Sf9 cells. The virus-containing supernatant was used to infect fresh Sf9 cells for 72 hrs and procedure was repeated twice to amplify the baculovirus. 20 μl of the third cycle supernatant was added to Sf9 cells in 250 ml SFM900-II medium (Invitrogen) with 5% FBS to express 6xHis-GFP-Dm-Kat60 at a low level (high levels of expression yielded inactive aggregates). Infected cells were lysed with a French press, centrifuged, and the resulting supernatant incubated with 1ml nickel resin (Qiagen) at 4 °C for 1 hr. The resin was then transferred into 20ml chromatography column (Biorad) and washed with 4 column volumes of wash buffer (40 mM imidazole in resuspension buffer without PMSF), and eluted with 500 mM imidazole in resuspension buffer without PMSF. The eluted rDm-Kat60 was concentrated by centrifugation and exchanged into severing buffer I (20 mM HEPES, pH 7.0, 300 mM NaCl, 3 mM MgCl₂, 10% sucrose, 50 μM ATP, 5 mM DTT).

In vitro assays

Rhodamine-labeled, taxol-stabilized non-polarity or polarity-marked MTs were prepared according to an online protocol (http://mitchison.med.harvard.edu/protocols.html) and immobilized in the flow chamber with a kinesin rigor mutant (G234A) as reported⁴⁰. The assay was conducted in severing buffer II (20 mM HEPES, pH 7.0, 100 mM NaCl, 3 mM MgCl₂, 10% sucrose, 2 mM ATP, 10 mM DTT, 7.5 mg/ml BSA, 0.1% Pluronic F-127) at 24 °C. An oxygen-scavenger system (220 μg/ml glucose oxidase, 22.5 mM glucose, 36 μg/ml catalase) was included to minimize photodamage. In some control assays, 5 mM AMP-PNP or 4 mM hexokinase was used in place of ATP. Subtilisin-pretreatment of MTs was the same as reported⁷. After perfusing MTs into the flow chamber cell, the purified rDm-Kat60 construct was introduced into the chamber and fluorescent MTs were imaged by Total internal reflection microscopy (TIRF). Our TIRF microscope used a home-made laser system assembled around a Nikon Eclipse Ti microscope and a high numerical aperture objective (60x, NA=1.49).

Digital images were saved as stacks of TIFF files, and time-lapse series were saved as AVI videos. Kymographs were generated using the Multiple Kymograph plug-in (ImageJ). From the kymographs of all MTs recorded for each treatment of the depolymerization assay, we calculated the average angle of the MT ends’ displacements through time. The depolymerization rate was determined by calculating the reciprocal of the tangent of the average angle, and then applying the appropriate conversions to derive the distance and time values. The in vitro severing frequency was determined by manually counting the number of severing events in every MT. The number of severing events in a single movie was divided by the total length of the MTs at the start of the movie and divided by the total time of the movie.

D17 cell RNAi and migration assays

D17 cells were maintained in a RT incubator in D17 medium: Schneider’s medium (Invitrogen) supplemented with 10% FBS (Gibco), 1% antibiotic-antimycotic (Gibco), and 1.25 μg/mL insulin (Invitrogen).

For RNAi, D17 cells were treated on days 0, 2, 4, 6, and 8 with 20 μg of dsRNA in 1 mL serum-containing Schneider cell medium (Life Technologies). On day 9, the RNAi-treated cells were either plated on ECM-coated plastic tissue culture dishes for wound-healing experiments or ECM-coated glass-bottom tissue culture dishes for single-cell analysis. For wound-healing experiments, cells were allowed to adhere for 4–6 hr before manually scraping the cell monolayer with a pipette tip. After replenishing the medium, the scratch-wound regions were imaged 0 hr and 24 hr after wounding using a phase-contrast microscope. For single-cell analysis, cells were imaged every 3 min for 3 hr using a phase-contrast microscope equipped with a 10x (0.25 NA) objective.

Electron microscopy

Samples for negative stain electron microscope experiments were prepared by mixing 1.5 μM of MTs with 50 nM rDm-Kat60 in severing buffer II (above) for 30 min at 24 °C. 5 μl of this mixture was then loaded onto freshly glow-discharged carbon supported grids (Electron Microscopy Sciences, Hatfield, PA). The grid was washed and stained with 1% uranyl acetate, followed by observation in a Tecnai F20 microscope (FEI, Eindhoven, The Netherlands) operating at 120 kV with a nominal magnification of x50,000.

Statistics

Differences between treatments were analyzed using either a one-way nonparametric analysis of variance (Kruskal-Wallis) for multiple group comparisons or a nonparametric t test (Mann-Whitney) for two group comparisons (SigmaStat, Systat Software; or GraphPad Prism, GraphPad Software). Measurement means were taken to be statistically different if P < 0.05.

Supplementary Material

Supplementary Figures

Supplemental Figure S1: Dm-Kat60’s cortical staining is reduced/abolished by Dm-Kat60 RNAi

(a) Comparison of Dm-Kat60 immunofluorescence in a field of control vs. Dm-Kat60 RNAi-treated S2 cells. Dm-Kat60’s cortical staining pattern appears to be specific as it was significantly reduced or abolished by the targeted depletion of Dm-Kat60 by RNA-mediated interference. On average, Kat60 RNAi in S2 cells reduced the intensity of cortical Dm-Kat60 immunofluorescence by 71% relative to controls (100 randomly selected cells measured in each condition; p<0.001). Moreover, ~30% (31/100) of the RNAi-treated cells displayed no detectable Dm-Kat60 staining at the cortex compared to only 2% (2/100) of controls showing the same.

(b) A Dm-Kat60 RNAi treated S2 cell double-labeled for Dm-Kat60 (green) and actin (red) further demonstrating that RNAi reduces the cortical staining of Dm-Kat60 but does not displace cortical actin.

Scale bar, 10 μm (panels a and b).

Supplemental Figure S2: Ectopically expressed GFP-Dm-Kat60 localizes to the cortex of interphase S2 cells and rescues the Dm-Kat60 RNAi phenotype.

(a & b) The cortical localization of Dm-Kat60 was confirmed in S2 cells expressing a Dm-Kat60-GFP fusion protein. Because overexpression of this construct is highly toxic to the cell, these experiments were performed in S2 cells depleted of endogenous Dm-Kat60 using dsRNA targeting the 3′ untranslated region (UTR) of the Dm-Kat60 mRNA. RNAi-treated cells were then transfected with an inducible plasmid encoding Dm-Kat60 (lacking the endogenous 3′UTR) fused to GFP, which was expressed at a level approximating that of the endogenous protein. Panel a is a Western blot indicating endogenous Dm-Kat60 and Dm-Kat60-GFP expression levels following the indicated treatments. The lower anti-actin blot is included as a loading control (see also Supplemental information, Fig. s8a). Panel B shows shows the typical localization of Dm-Kat60-GFP (labeled with an anti-GFP antibody) and MTs (labeled with an anti-α-tubulin antibody) in a transfected interphase S2 cell. Far right panel (**) is a high magnification of the boxed region of the cell in the merged panel. Indirect immunofluorescence was used to localize GFP-Dm-Kat60 because the GFP signal alone was too low for the characterization of any specific localization pattern. Clearly apparent cortical staining was observed in ~67% (311/469) of Dm-Kat60-GFP transfected cells as compared to <5% (11/521) of non-transfected cells stained with the same antibody.

(c & d) Replacement of endogenous Dm-Kat60 with the GFP-tagged fusion protein rescued the Dm-Kat60 RNAi phenotype. Panel C shows Immunofluorescence micrographs indicating the typical organization of MT arrays in control, Dm-Kat60 RNAi, and Dm-Kat60 RNAi/Dm-Kat60-GFP transfected interphase S2 cells. (d) Most Dm-Kat60 depleted cells have dense arrays of curled MTs arranged parallel to the cortex, but this phenotype is rescued once Dm-Kat60 RNAi cells are induced to express RNAi resistant Dm-Kat60-GFP. Rescue requires catalytically active Dm-Kat60, as demonstrated by the inability of the inactive mutant, Dm-Kat60 (E393Q)⁴¹, to restore a normal cortical MT arrangement. p = 0.0216. Data represent Mean ± SEM from three independent experiments.

Scale bars, 10 μm (panels a and b).

Supplemental Figure S3: The cortical localization of Dm-Kat60 does not require MTs but is reliant upon actin.

(a) Confocal micrographs showing that the localization of Dm-Kat60 at the cortex persists after microtubule disassembly with colchicine.

(b) Cortical Dm-Kat-60 staining is lost when actin is disassembled by cytochalasin D.

Scale bar, 10 μm (panels a,b).

Supplemental Figure S4: Comparison of MT plus-end trajectories generated using our automated algorithm vs. hand tracking.

The image shows an individual time-point from a live cell movie of MT behaviors in a control GFP-α-tubulin-expressing interphase S2 cell. The trajectory paths of MT ends generated automatically are shown in magenta while the hand-tracked trajectory paths of these same MT ends are shown in blue. Scale bar, 5μm. In all, the automated algorithm correctly identified and followed the movement 98.3% of MT plus-ends (n=290).

Supplemental Figure S5: MT severing enzymes in D17 cells.

(a) Immunolocalization of Spastin, Fidgetin, and tubulin in Drosophila D17 cells. The boxed areas are magnified in the insets.

(b) Quantification of migration during wound closure for control, Spastin, and Fidgetin RNAi-treated D17 cells. Migration area was calculated by subtracting the total wound area at 24 hr from the total wound area at 0 hr after wounding and normalized to control RNAi.

(c) Immunolocalization of actin in control and Dm-Kat60 RNAi-treated D17 cells. Scale bars, 10 μm for both panel a and c

Supplemental Figure S6: Human KATNA1 localizes to the leading edge of migratory cells and its depletion by RNAi affects microtubule organization and cell migration similarly to Dm-Kat60.

(a) Immunofluorescence of human HS578T cells stained with our previously characterized anti-KATNA1 antibody³⁵ revealed that, similarly to Dm-Kat60, KATNA1 adopts a cortical localization that is most prominent at the leading edge of polarized cells as well as the edge of additional cellular protrusions (arrow). The area of the leading edge showing pronounced KATNA1 staining boxed in the merged panel is magnified in the inset.

(b) SiRNA depletion of KATNA1 induces a dramatic alteration in cell morphology hallmarked by an increase in the length of cellular protrusions which filled with densely packed arrays of parallel MT. The image in this panel shows representative fields of control and KATNA1 siRNA-treated Hs578T cells immunostained for α-tubulin.

(c) SiRNA depletion of KATNA1 increases the average length of the cellular protrusions more than 2-fold. p< 0.0001. Data represent mean ± SEM. Numbers in the columns indicate the number of protrusions measured/total number of cells analyzed.

(d) KATNA1 siRNA treated cells migrated ~1.5 fold faster than controls in wound healing assays. Monolayers of control and KATNA1 siRNA-treated Hs578T cells were wounded and imaged by phase microscopy immediately and 14 hrs later. The relative positions of the wound edge at the beginning and end of the experiment are indicated by green and yellow lines, respectively.

(e) Quantification of the rate of wound closure in control vs. KATNA1 siRNA treated cultures The data represent the mean ± SEM of three independent experiments. p = 0.0328

Scale bar, 20 μm (panel a) or 100 μm (panels b, d).

Supplemental Figure S7: Model of kat-p60 activity at the cortex of interphase cells.Dm-Kat60 localizes to the cortex of lamellipodia (right), possibly by interacting with the F-actin enriched in this region. Our results suggest that as plus-ends of growing MTs approach the cortex, Dm-Kat60 attacks both the ends and the walls of MTs and perhaps eliminates stabilizing EB1 “caps” at plus-ends. Following Dm-Kat60 induced catastrophe and/or depolymerization at the cortex, kinesin-13 may assume responsibility for plus-end disassembly. If Dm-Kat60 is depleted (left), the loss of this potent catastrophe factor increases the lifetimes of MTs invading the cortex. Consequently, the MTs continue to grow even after reaching the cell boundary, and so the cell periphery fills with dense arrays of MTs running parallel to the cortex. MTs within these arrays could conceivably lose the ability to respond normally to the intra- and extra-cellular cues that optimize their dynamics and organization to stimulate movement, morphogenesis, etc …

Supplemental Figure S8. Electrophoretic data in uncropped form.

Panel a shows the scanned image of the Western blot for Supplemental Figure S2a. To avoid cross reaction of different antibodies, the transferred membrane was cut into two parts and developed separately for Dm-kat60/Dm-kat60-GFP or actin (arrows). Panel b shows the scanned image of the original of the original Western blot in Figure 4b. Dm-Kat60 and actin was detected at the same time (arrows). Panel c shows scanned image of the original SDS-PAGE of purified rDm-kat60 used in Figure 7a. Molecular markers are shown for each panel.

Supplemental Movie S1: a &b: Time-series movie of MT plus-end dynamics near the cortex of a control (a) or Dm-Kat60 RNAi (b) treated GFP-tubulin expressing interphase S2 cell. Each image is a maximum intensity projection of three 1 μm optical sections; image stacks were collected at 5 second intervals. The cell was imaged using a 100X, 1.4 NA objective. c: Time-series movie showing severing near the plus-end of a MT at the cortex in a control treated GFP-α-tubulin expressing interphase S2 cell. Each image is a maximum intensity projection of three 1 μm optical sections and image stacks were acquired at three second intervals. d & e: Movie showing MT plus-ends tracked with our automated algorith in control (d) or Dm-Kat60 RNAi (e) treated S2 cells. Growing ends are marked with blue, shrinking ends with pink and pausing ends with yellow dots. The blue tail shows the last few frames of the trajectory. The time between frames is 5 seconds.

Supplemental Movie S2: Time-series movies showing the motility of individual control (left) and Dm-Kat60 RNAi-treated (right) D17 cells imaged with a 10x phase contrast objective. 4 trajectories of individual cell movements are highlighted by colored lines in each field. Total time of imaging is 3 hours and time interval is 3 mins.

Supplemental Movie S3: Time series movies showing representative cortical zones of S2 cells transfected with GFP-tubulin and mcherry-actin after control (left) or Dm-Kat60 RNAi (right) treatments. Cells were imaged with a 100X, 1.4 NA objective at 10 second intervals.

Supplemental Movie S4: Time series of TIRF images showing rhodamine-labeled MTs fixed on a coverslip and incubated with 50 nM GFP-Dm-Kat60. Time in seconds is shown in the upper left of each image. Negative numbers indicate time before the introduction of Dm-Kat60 and positive numbers indicate time after the addition of Dm-Kat60. The movie is paused at t=0.