Many cell types contain a subset of long-lived, ‘stable’ microtubules that differ from dynamic microtubules in that they are enriched in post-translationally detyrosinated tubulin (Glu-tubulin). Elevated Glu tubulin does not stabilize the microtubules and the mechanism for the stability of Glu microtubules is not known. We used detergent-extracted cell models to investigate the nature of Glu microtubule stability. In these cell models, Glu microtubules did not incorporate exogenously added tubulin subunits on their distal ends, while >70% of the bulk microtubules did. Ca2+-generated fragments of Glu microtubules incorporated tubulin, showing that Glu microtubule ends are capped. Consistent with this, Glu microtubules in cell models were resistant to dilution-induced breakdown. Known microtubule end-associated proteins (EB1, APC, p150Glued and vinculin focal adhesions) were not localized on Glu microtubule ends. ATP, but not nonhydrolyzable analogues, induced depolymerization of Glu microtubules in cell models. Timelapse and photobleaching studies showed that ATP triggered subunit loss from the plus end. ATP breakdown of Glu microtubules was inhibited by AMP-PNP and vanadate, but not by kinase or other inhibitors. Additional experiments showed that conventional kinesin or kif3 were not involved in Glu microtubule capping. We conclude that Glu microtubules are stabilized by a plus-end cap that includes an ATPase with properties similar to kinesins.

Microtubules (MTs) function in mitosis, cell motility, intracellular transport and the maintenance of cell shape. At least two populations of MTs have been distinguished in nonmitotic cells: short-lived or dynamic MTs (t1/2=5-10 minutes) and long-lived or stable MTs (t1/2>1 hour) (Schulze and Kirschner, 1987; Webster et al., 1987a). Dynamic MTs predominate in proliferating and undifferentiated cells, whereas stable MTs may be the more abundant type in differentiated cells (Gundersen and Bulinski, 1986; Bulinski and Gundersen, 1991).

An important question concerning stable MTs is what gives rise to their heightened stability, while adjacent MTs remain dynamic. In many cell types, stable MTs accumulate a post-translationally modified form of tubulin known as detyrosinated tubulin (or Glu-tubulin), whereas dynamic MTs contain predominantly tyrosinated tubulin (Tyr-tubulin) (reviewed by Bulinski and Gundersen, 1991; Liao et al., 1999). MTs that accumulate sufficient Glu-tubulin to be detected by immunofluorescence with antibody to Glu tubulin (Glu MTs) have been shown to be stable MTs, whereas those that stain with an antibody to Tyr-tubulin, but not with an antibody to Glu-tubulin (Tyr MTs), have been shown to be dynamic MTs (Gundersen et al., 1984; Gundersen et al., 1987a; Kreis, 1987; Khawaja et al., 1988; Schulze et al., 1987; Webster et al., 1987a). Despite the correlation between elevated levels of Glu-tubulin and MT stability, detyrosination does not directly cause MT stability (Khawaja et al., 1988; Webster et al., 1990; Cook et al., 1998). Instead, as stabilized MTs persist in the cell, they become increasingly detyrosinated (Gundersen et al., 1987a).

MT-associated proteins (MAPs) are candidates for the selective stabilization of MTs. Well-characterized MAPs (MAP1, MAP2, MAP4 and tau) bind along the length of MTs and increase their stability (Hirokawa, 1994). However, none of these proteins has been shown to generate MT subsets of different stabilities within a single cell. When present, these MAPs enhance the stability of all cellular MTs (Hirokawa, 1994). Consistent with this, the distribution of MAP4 on Tyr and Glu MTs showed no evidence of segregation and MAP4 does not exhibit increased binding to Glu MTs (Chapin and Bulinski, 1994). Other proteins known to stabilize MTs, e.g. STOP proteins (Margolis et al., 1986), also do not appear to selectively stabilize MTs.

The selective stabilization of Glu MTs may involve a unique mechanism. Glu MTs are refractory to subunit addition and loss in vivo, suggesting that these MTs may be capped (Gundersen et al., 1987b; Webster et al., 1987a; Schulze and Kirschner, 1987). Consistent with this, Glu MTs are resistant to end-mediated depolymerization by nocodazole or by dilution (Khawaja et al., 1988). Also, protein phosphatase inhibitors cause the selective breakdown of Glu MTs, with most Tyr MTs remaining intact (Gurland and Gundersen, 1993; Merrick et al., 1997). This suggests that stable MTs are regulated differently from dynamic MTs by protein phosphorylation.

To understand how Glu MTs are stabilized, we developed an extracted cell model that preserves the stability and assembly refractoriness of Glu MTs. With these models, we show that a functional cap exists on the plus ends of Glu MTs. Known MT end-binding proteins were not detected on Glu MT plus ends. We show that the stability of the Glu MTs in the extracted cell system can be modified by ATP hydrolysis. By timelapse and photobleaching studies, we found that ATP hydrolysis triggers Glu MTs depolymerization from the plus end, suggesting that the ATPase is part of the cap. Nucleotide and inhibitor profiles suggest that the ATP effect is not due to the action of a kinase, but may be mediated by a kinesin-like protein. These results provide the first evidence that the selective stabilization of Glu MTs in vivo occurs via a capping mechanism.

Reagents

Reagents used in this study were purchased from Sigma Chemical (St Louis, MO, USA), except where noted. UTP was from Pharmacia (Piscataway, NJ, USA), lavendustin was from Gibco, Pipes was from Research Organics (Cleveland, OH, USA) or Sigma, and taxol was a gift from the National Cancer Institute.

Cell culture and preparation of extracted cell models

African green monkey kidney epithelial cells (TC-7) were cultured in DMEM (Gibco, Grand Island, NY, USA) with 10% FBS (Gibco) and 20 mM Hepes as previously described (Gundersen et al., 1984). Cells plated on acid-washed sterile coverslips were used after 2 days when they were approximately 50% confluent. To prepare extracted cell models, cells were rinsed twice in PEM (100 mM Pipes, pH 6.9, 1 mM EGTA, 2 mM MgCl2) and extracted with 0.2% Triton X-100 in PEM for 1 minute at 37°C. Extraction with saponin (200 μg/ml for 1 minute) gave similar results. Extracted cells were rinsed with PEM, and then incubated with tubulin subunits to test for MT growth or treated to test MT stability. In some experiments, cells were treated with 10 μM nocodazole for 4 hours before extraction and growth.

MT growth assay

Immediately after extraction, cells on coverslips were inverted onto 50 μl of tubulin solution (purified calf brain tubulin in PEM containing 1 mM GTP) on parafilm and incubated at 37°C for up to 5 minutes. Tubulin was purified by cycles of assembly/disassembly followed by DEAE-Sephadex chromatography as described (Mikhailov and Gundersen, 1995). The tubulin concentration in the incubations was varied between 0.03 mg/ml and 0.5 mg/ml. Tubulin concentrations >1 mg/ml could not be used, since the tubulin began to self-assemble. In some experiments, we included 0.2 mg/ml casein (dialyzed exhaustively against PEM) in the incubations to reduce nonspecific absorption of tubulin. Coverslips were fixed in −20°C methanol.

After triple immunofluorescence staining for Glu, Tyr and brain βIII tubulin (see below), samples were examined with a Nikon PlanApo 100× lens on a Nikon Optiphot (see Gurland and Gundersen, 1995). Images were captured with an SIT camera (Dage-MTI, Michigan City, IN, USA) (32-frame average) and processed with Image-1 software. The three images (Glu, Tyr and βIII tubulin) were aligned and then an on-screen overlay was used to mark the ends of Glu and Tyr MTs; this overlay was placed on the image of the βIII tubulin segments to identify growing MT ends.

Ca2+ fragmentation of MTs

Extracted cells were treated with 0.8-2 mM CaCl2 in PEM for 0.5-5 minutes to fragment Glu MTs. This yielded free Ca2+ concentrations of 1 μM-1 mM, as determined by a Ca2+ selective electrode (>10 μM), and by a computer program (<10 μM) (courtesy of P. Brandt, Columbia University). Ca2+-treated cytoskeletons were incubated with tubulin and processed as above.

MT depolymerization assay

Extracted cells were treated with nucleotides or pharmacological inhibitors in order to test the stability of Glu MTs. Extracted cells were incubated for 60 minutes in 1 ml of PEM or inverted on 50 μl of PEM on parafilm (similar results were obtained with both techniques). Drugs were dissolved directly in PEM or in DMSO stock solutions that were added to PEM such that the final DMSO concentration did not exceed 0.1% (v/v). DMSO at 0.1% had no effect on MTs in the extracted cells. All solutions and incubations were at 37°C. Following treatments, extracted cells were rinsed in PEM and fixed in −20°C methanol.

Timelapse recordings of fluorescent MTs

TC-7 cells were pressure-microinjected with 4 mg/ml rhodamine-tubulin (R-tubulin). R-tubulin was prepared as described (Mikhailov and Gundersen, 1995). For experiments with extracted cells, R-tubulin-injected cells were incubated overnight to ensure that R-tubulin had incorporated into stable MTs. For experiments on living cells, R-tubulin was allowed to incorporate for 2-3 hours, which was sufficient to label the dynamic MTs.

Live cell recordings

Coverslips were mounted in a Sykes-Moore chamber (Bellco Glass Co., Vineland NJ, USA) and imaged under reduced oxygen conditions using Oxyrase, as described (Mikhailov and Gundersen, 1995). Images were acquired every 5 seconds. Throughout the recording, cells were perfused at a rate of 0.1 ml/minute with medium treated with Oxyrase to reduce photodamage (Mikhailov and Gundersen, 1995).

Extracted cell recordings

Coverslips containing injected cells, were assembled into a Sykes-Moore perfusion chamber (0.75 ml) and perfused with Oxyrase-treated medium. Cells were extracted on the microscope by perfusing (0.5 ml/minute) with 1 ml each of medium, PEM, then 0.2% Triton X-100 in PEM. All solutions were at 37°C. PEM containing 40 mM dl-lactic acid and 1:50 Oxyrase was then perfused at 0.5 ml/minute for 2 minutes, then at 0.1 ml/minute for 15 minutes. Lactic acid was included as a source of reducing sugar for the Oxyrase (Oxyrase brochure). The 15 minute post-extraction incubation allowed dynamic MTs to depolymerize, leaving only stable MTs; this was confirmed by immunolabelling with Glu-tubulin antibody. For ATP treatment, we perfused extracted cells with 200 μM ATP in PEM, lactic acid and Oxyrase at 1 ml/minute for 1 minute and then 0.2 ml/minute for the remainder of the experiment.

Depolymerization rates of MTs in recordings of live and extracted cells were measured by tracing MT lengths using the ‘trace curve length’ function on Image-1. Coordinates of the traces were transferred to a spread sheet program, which derived the depolymerization rates (Mikhailov and Gundersen, 1998). Data was obtained from images acquired every 5 seconds.

Photobleaching experiments

Extracted cells with R-labeled, stable MTs were prepared as described above, except that in some experiments, dynamic MTs were allowed to depolymerize for up to 30 minutes before perfusing with ATP. Extracted cells in Sykes-Moore chambers were imaged with a 63× Planapo Chromat and a Zeiss IM-35 inverted microscope. MTs were exposed to a 200 mW pulse of 514 nm laser radiation for 50 milliseconds, focused into a bar with a cylindrical lens. The photobleaching apparatus has been described in detail previously (Gorbsky et al., 1987; Rodionov et al., 1994). This exposure permanently inactivated rhodamine fluorescence at fixed sites on the MTs, without damaging the MTs (see text). After photobleaching, ATP was perfused as described above. Images were recorded every 10 seconds with a Photometrics cooled CCD (512×512 pixel Tektronix chip) (Tucson, AZ, USA) and processed with MetaMorph software (Universal Imaging, West Chester PA, USA). We usually took 10 images before photobleaching and ATP addition to establish an unstimulated rate of MT depolymerization.

Indirect immunofluorescence

Fixed preparations were stained by triple indirect immunofluorescence with a rat antibody specific for Tyr tubulin (YL 1/2) (Kilmartin et al., 1982), a rabbit antibody specific for Glu tubulin (SG) (Gundersen et al., 1984), and various mouse monoclonal antibodies. YL 1/2 hybridoma was purchased from the European Collection of Animal Cell Cultures (Salisbury, UK). Secondary antibodies from Jackson ImmunoResearch (West Grove, PA, USA) were used as previously described (Gurland and Gundersen, 1995). For MT growth experiments, we used a mouse monoclonal antibody specific for the βIII brain-specific isoform of β-tubulin (Banerjee et al., 1990). βIII antibody was the generous gift of R. Ludueña (U. Texas, San Antonio, USA). We amplified the βIII signal by using biotinylated goat anti-mouse IgG and then rhodamine-conjugated streptavidin (both from Jackson ImmunoResearch).

For localization of proteins on the ends of MTs, we used the following mouse antibodies: SUK4 kinesin antibody (Ingold et al., 1988) (Developmental Studies Hybridoma Bank), EB1 antibody (Clone 5, Transduction Laboratories, Lexington, KY, USA) and vinculin antibody (Clone VIN 11-5, Sigma). We used a guinea pig antibody to Glu tubulin in experiments in which we localized APC and p150Glued with rabbit antibodies. The guinea pig antibody is specific for Glu tubulin by western blotting, and stains a similar subset of MTs in TC-7 cells as the rabbit SG antibody (data not shown). Rabbit APC antibody (Nathke et al., 1996) was a gift from I. Nathke (Stanford University) and rabbit p150Glued antibody (d’Art; Vaughan et al., 1999) was a gift from R. Vallee (UMass-Worchester). Preparations were imaged with a cooled CCD camera with a Kodak KAF 1400 chip (1317×1035 pixels) (MicroMAX; Princeton Instruments, Trenton, NJ, USA) on a Nikon Optiphot microscope. Images were processed with Metamorph software.

Protein phosphorylation of extracted cells

TC-7 cells plated in 35 mm dishes were grown to 75-90% confluency, extracted (as above) and then incubated in 0.5 ml labeling buffer (50 mM β-glycerophosphate, 50 mM Pipes, pH 6.9, 1 mM EGTA, 2 mM MgCl 2, 0.2 mM PMSF, 0.1 μg/ml CLAP (chymostatin, leupeptin, antipain, pepstatin), 5 mM NaF, 1 μM Calyculin A) with 0.1 mM ATP and 5 μCi [γ-32 P]ATP (Amersham) per dish. Inhibitors were added at the same time as [γ-32 P]ATP. After 1 hour, the soluble fraction was collected and TCA-precipitated. The remaining cytoskeletal fraction was rinsed with PEM and solubilized in SDS-sample buffer. Soluble and cytoskeletal fractions from equal numbers of cells were subjected to SDS-PAGE and autoradiography.

Tyr MTs in extracted cytoskeletons are growth-competent

Since intact cellular Glu MTs cannot be isolated by subcellular fractionation, we developed a detergent-extracted cell model to explore the mechanism of Glu MT stability. From earlier work (e.g. Khawaja et al., 1988), we knew that Glu MTs were resistant to dilution-induced depolymerization after cell extraction, but it was important to test whether they also retained their in vivo characteristic of resistance to tubulin addition (Webster et al., 1987a).

To examine this, we added purified brain tubulin to extracted cells and then detected the added tubulin with an antibody to βIII tubulin, a minor brain-specific isoform (Banerjee et al., 1990). In extracted cells incubated with tubulin, segments of brain tubulin were found throughout the cell and were aligned radially (Fig. 1b), as were the endogenous MTs (Fig. 1a). The segments were at the ends of existing Tyr MTs rather than free in the cytoskeleton, demonstrating that many Tyr MTs have elongated with the added tubulin (Fig. 1c,d). Segments were not detected in cell-free areas. If MTs were depolymerized before cell extraction (Fig. 1e), the added tubulin only grew from the centrosome (Fig. 1f). Thus, added brain tubulin grows from pre-existing MTs and the centrosome in extracted cells.

Fig. 1.

Exogenous bovine brain tubulin adds to the distal ends of MTs in extracted TC-7 cells. Shown are immunofluorescent images of cells stained for Tyr tubulin (a,c,e) and brain βIII tubulin (b,d,f). (a,b) Low magnification showing the radial distribution of added brain tubulin segments. (c,d) Higher magnification showing added brain segments at the ends of endogenous MTs. Arrows indicate ends of some of the growing Tyr MTs. (e,f) Cells treated with 10 μM nocodazole (4 hours) to depolymerize all the MTs before extraction. (e) No incubation with brain tubulin, showing depolymerization of all MTs by nocodazole. (f) Incubation with brain tubulin, showing growth of MTs from centrosomes. Bars, 5 μm.

Fig. 1.

Exogenous bovine brain tubulin adds to the distal ends of MTs in extracted TC-7 cells. Shown are immunofluorescent images of cells stained for Tyr tubulin (a,c,e) and brain βIII tubulin (b,d,f). (a,b) Low magnification showing the radial distribution of added brain tubulin segments. (c,d) Higher magnification showing added brain segments at the ends of endogenous MTs. Arrows indicate ends of some of the growing Tyr MTs. (e,f) Cells treated with 10 μM nocodazole (4 hours) to depolymerize all the MTs before extraction. (e) No incubation with brain tubulin, showing depolymerization of all MTs by nocodazole. (f) Incubation with brain tubulin, showing growth of MTs from centrosomes. Bars, 5 μm.

We optimized the growth conditions by varying the tubulin concentration, incubation times, nucleotide composition and stabilizing factors. Growth was maximal at 0.1 mg/ml tubulin (Fig. 2); above this concentration the percentage of growing MTs did not increase significantly, and it became difficult to detect the exogenous tubulin because of higher background staining with the βIII antibody. Incubation time was not critical, but 2 minutes gave a reasonable segment length (approx. 5 μm). No growth was observed in the absence of GTP. Addition of 5 μM taxol, which stabilizes MTs (Shiff and Horowitz, 1980), increased the fraction of growing MTs to 70% (Fig. 2). This is very near the 80% of MTs that incorporate segments at their ends during a 2 minute pulse in vivo (Schulze and Kirschner, 1987).

Fig. 2.

Optimization of growing MTs in extracted cells. Cells were extracted and incubated with the indicated concentrations of brain tubulin with and without 5 μM taxol. For each condition, >180 MT ends were counted (data were pooled from three experiments, which gave similar results).

Fig. 2.

Optimization of growing MTs in extracted cells. Cells were extracted and incubated with the indicated concentrations of brain tubulin with and without 5 μM taxol. For each condition, >180 MT ends were counted (data were pooled from three experiments, which gave similar results).

Glu MTs in extracted cytoskeletons are not growth-competent

Using optimal conditions for growth (0.1 mg/ml tubulin, 5 μM taxol, 1 mM GTP, for 2 minutes), we examined whether Glu MTs were able to incorporate tubulin. We found that Glu MTs did not grow, even though Tyr MTs in the same part of the cell did (Fig. 3a). In six experiments in which over 600 Glu MTs were examined, we detected only one Glu MT that appeared to grow. Thus, Glu MTs in our extracted cell system retain the refractoriness to tubulin subunit addition that they have in living cells.

Fig. 3.

Brain tubulin adds to Tyr MTs but not to Glu MTs, unless Glu MTs are first fragmented by calcium treatment. (a) Extracted cells incubated with brain tubulin: blue, Tyr tubulin; green, Glu tubulin; and red, brain βIII tubulin. Many growing Tyr MTs are observed (red segments on ends of blue MTs). Arrows point to Glu MTs (green) that do not have brain tubulin segments (red) on their ends. (b) Extracted cells treated with calcium to fragment Glu MTs before incubating with brain tubulin. Arrows indicate Glu MT fragments (green) extended by brain tubulin segments (red). Bars, 5 μm.

Fig. 3.

Brain tubulin adds to Tyr MTs but not to Glu MTs, unless Glu MTs are first fragmented by calcium treatment. (a) Extracted cells incubated with brain tubulin: blue, Tyr tubulin; green, Glu tubulin; and red, brain βIII tubulin. Many growing Tyr MTs are observed (red segments on ends of blue MTs). Arrows point to Glu MTs (green) that do not have brain tubulin segments (red) on their ends. (b) Extracted cells treated with calcium to fragment Glu MTs before incubating with brain tubulin. Arrows indicate Glu MT fragments (green) extended by brain tubulin segments (red). Bars, 5 μm.

Glu MTs can grow if internal sites are exposed by fragmentation

The growth resistance of Glu MTs may be due to factors acting on the ends of MTs or along the length of MTs. To distinguish between these possibilities, we treated Glu MTs in extracted cells with Ca2+, which is known to fragment MTs (Schliwa et al., 1981). We confirmed by recordings of R-tubulin-labeled MTs in extracted cells that >5 μM free Ca 2+ caused infrequent breaks along the length of stable MTs (data not shown). Brain tubulin added to calcium-treated, extracted cells with fragmented Glu MT, consistently grew off one end of many Glu MT segments (Fig. 3b). Growth was almost always from only one end, which is probably the plus end, since plus-end growth is favored by the low tubulin concentration and short incubation we used. These results demonstrate that internal sites on Glu MTs can grow if exposed and strongly suggest that factors are acting on the ends of Glu MTs to prevent growth.

Glu MTs are more resistant to dilution than Tyr MTs

An earlier study showed that Glu MTs are more resistant to dilution than Tyr MTs in TC-7 cells (Khawaja et al., 1988) and we confirmed this result in the present study. Intact TC-7 cells contain many more Tyr MTs than Glu MTs (Fig. 4a,b). In contrast, extracted TC-7 cells that were incubated in PEM buffer for 15 minutes contained similar levels of Tyr and Glu MTs (Fig. 4c,d), showing that many Tyr MTs, but few Glu MTs, were depolymerized by dilution within 15 minutes. By 60 minutes incubation in PEM, virtually all remaining MTs were Glu MTs (Fig. 4e,f). Note that much of the Tyr tubulin staining at 60 minutes is due to weak staining of the Glu MTs by Tyr antibody. Most Glu MTs were resistant to dilution for up to 60 minutes (compare Fig. 4b and f) and many were present at 90 minutes, the longest time at which we looked (not shown).

Fig. 4.

Glu MTs are more resistant to dilution than Tyr MTs. Extracted TC-7 cells incubated in PEM for 0 minutes (a,b), 15 minutes (c,d) and 60 minutes (e,f). Shown are immunofluorescent images of cells stained with antibodies specific for Tyr-(a,c,e) and Glu-tubulin (b,d,f). Bar, 20 μm.

Fig. 4.

Glu MTs are more resistant to dilution than Tyr MTs. Extracted TC-7 cells incubated in PEM for 0 minutes (a,b), 15 minutes (c,d) and 60 minutes (e,f). Shown are immunofluorescent images of cells stained with antibodies specific for Tyr-(a,c,e) and Glu-tubulin (b,d,f). Bar, 20 μm.

As shown above (Fig. 3), Ca 2+ -treated Glu MT fragments were able to incorporate added tubulin. If tubulin was not added to these fragmented Glu MTs, they depolymerized within 10 minutes (not shown). We timelapse recorded the calcium-induced depolymerization of Glu MTs and directly observed depolymerization of the fragmented stable MTs from newly created plus and minus ends (data not shown). These results show that the extracted cell models used here faithfully maintain both growth refractoriness and stability of Glu MTs and support the idea that Glu MTs are capped.

MT end-binding proteins do not cap Glu MTs

There are a number of proteins that have recently been localized at or near the ends of MTs in vivo (see Table 1). We examined the possibility that one or more of these proteins might be capping Glu MTs by colocalizing these proteins with Glu and Tyr MTs in TC-7 cells. EB1 is an APC binding protein that is preferentially localized on the ends of MTs in cells (Berrueta et al., 1998; Morrison et al., 1998). We found EB1 spots on the ends of many Tyr MTs, but only on a few Glu MTs (Fig. 5). Analyses of aligned and overlaid images showed that 72% (n=309) of Tyr MT ends but only 13% of Glu MT ends (n=84) were positive for EB1. This suggests that EB1 localization on the ends of Glu MTs is not responsible for Glu MT stability. APC has been localized near the ends of MTs in cells (Nathke et al., 1996). We did not detect APC on the ends of Glu MTs, although we did observe APC accumulations near MT ends as previously reported (data not shown). MTs have been reported to be stabilized by interaction with focal adhesions or focal complexes (Kaverina et al., 1998). We colocalized Glu MTs with vinculin-stained focal adhesions and focal complexes and found that <10% of the Glu MT ends colocalized with these structures (Fig. 5g-i). Tyr MT ends were colocalized with large focal adhesions infrequently (<10%), but showed higher colocalization with small focal complexes (18%, n=261) (Fig. 5j-l). Others have found that Glu MT ends do not stain for p150Glued, a component of dynactin that is specifically localized at the ends of MTs (Vaughan et al., 1999), and we confirmed this in our experiments (data not shown). CLIP-170, another MT end binding protein, is specifically localized on growing MT ends, which would exclude its localization on the ends of Glu MTs (Perez et al., 1999). This lack of colocalization of MT-end-binding proteins with ends of Glu MTs suggests that the activity responsible for Glu MT capping is novel.

Table 1.

Localization of known MT end-associated proteins with Glu MT ends

Localization of known MT end-associated proteins with Glu MT ends
Localization of known MT end-associated proteins with Glu MT ends
Fig. 5.

Glu MT ends do not colocalize with EBI or vinculin. Unextracted TC-7 cells were stained for Glu tubulin (a,g), Tyr tubulin (d,j), and either EBI (b,e) or vinculin (h,k). (c) Overlay of a and b; (f) overlay of d and e; (i) overlay of g and h; (l) overlay of j and k. Bar, 5 μm.

Fig. 5.

Glu MT ends do not colocalize with EBI or vinculin. Unextracted TC-7 cells were stained for Glu tubulin (a,g), Tyr tubulin (d,j), and either EBI (b,e) or vinculin (h,k). (c) Overlay of a and b; (f) overlay of d and e; (i) overlay of g and h; (l) overlay of j and k. Bar, 5 μm.

ATP hydrolysis induces Glu MT breakdown

To characterize the Glu MT capping activity, we tested the ability of reagents to break down Glu MTs in extracted cytoskeletons. We suspected that ATP would induce Glu MT breakdown, since earlier work by Bershadsky and Gelfand (1981) showed that MTs in azide-treated cells broke down after extraction and ATP addition. This study did not distinguish between Tyr and Glu MTs. Additionally, Glu MTs, but not Tyr MTs are depolymerized by phosphatase inhibitors in many cells (Gurland and Gundersen, 1993; Merrick et al., 1997).

ATP treatment of the extracted cytoskeletons caused a concentration- and time-dependent loss of Glu MTs (Fig. 6). As little as 10 μM ATP caused detectable Glu MT breakdown and 100 μM ATP caused substantial Glu MT breakdown within 10 minutes (Fig. 6, Table 2). Over 90% of Glu MTs were disassembled in 60 minutes by 100 μM ATP (Fig. 6c). Fragments of MTs were occasionally observed after the ATP treatments; however, most MTs remaining were either full length or slightly shorter than untreated MTs. As Tyr MTs depolymerize in PEM alone and do not appear to be capped, we did not explore the effect of ATP on Tyr MTs.

Table 2.

Nucleotide-induced breakdown of Glu MTs

Nucleotide-induced breakdown of Glu MTs
Nucleotide-induced breakdown of Glu MTs
Fig. 6.

ATP hydrolysis induces Glu MT breakdown in extracted cells. In all panels, Glu tubulin immunofluorescence is shown. (a) Cells fixed immediately after extraction; (b,c) extracted cells treated with 0.1 mM ATP for 10 minutes (b) and 60 minutes (c); (d) extracted cells incubated with 1 mM AMP-PNP 60 minutes. Bar, 20 μm.

Fig. 6.

ATP hydrolysis induces Glu MT breakdown in extracted cells. In all panels, Glu tubulin immunofluorescence is shown. (a) Cells fixed immediately after extraction; (b,c) extracted cells treated with 0.1 mM ATP for 10 minutes (b) and 60 minutes (c); (d) extracted cells incubated with 1 mM AMP-PNP 60 minutes. Bar, 20 μm.

Glu MT breakdown occurred with other nucleotide triphosphates, but required higher concentrations (Table 2). ATP hydrolysis was required since ADP, AMP, PPi and the nonhydrolyzable analogs, AMP-PCP and AMP-PNP did not induce Glu MT breakdown (Fig. 6d, Table 2). These results suggest that an ATPase is responsible for the ATP-induced breakdown of Glu MTs.

Uncoupling of the ATP hydrolysis step from the disassembly of Glu MTs

To determine whether continuous ATP hydrolysis was necessary for the ATP-induced breakdown of Glu MTs, we sought conditions that might separate ATP hydrolysis from the depolymerization of Glu MTs. Promoters of MT-subunit interactions, such as glycerol and taxol, blocked ATP-induced breakdown of Glu MTs. Similar numbers of Glu MTs persisted in extracted cells treated with ATP in the presence of 20% glycerol (or 10 μM taxol) as in unextracted cells or unincubated extracted cells (compare Figs 7b and 4b). This suggests that ATP stimulated a typical end-dependent breakdown of the Glu MTs.

Fig. 7.

Glycerol reversibly blocks ATP-induced breakdown of Glu MTs. Glu tubulin immunofluorescence is shown in all panels. Extracted cells were incubated in 0.1 mM ATP in the absence (a) or presence of 20% glycerol for 60 minutes (b). (c) Extracted cells were incubated in 20% glycerol and 0.1 mM ATP for 30 minutes, rinsed and then further incubated in PEM for 60 minutes. (d) Extracted cells were incubated in 20% glycerol without ATP for 30 minutes, rinsed and then further incubated in PEM for 60 minutes. Bar, 20 μm.

Fig. 7.

Glycerol reversibly blocks ATP-induced breakdown of Glu MTs. Glu tubulin immunofluorescence is shown in all panels. Extracted cells were incubated in 0.1 mM ATP in the absence (a) or presence of 20% glycerol for 60 minutes (b). (c) Extracted cells were incubated in 20% glycerol and 0.1 mM ATP for 30 minutes, rinsed and then further incubated in PEM for 60 minutes. (d) Extracted cells were incubated in 20% glycerol without ATP for 30 minutes, rinsed and then further incubated in PEM for 60 minutes. Bar, 20 μm.

We next asked whether Glu MTs treated with ATP in the presence of glycerol would break down if the ATP and glycerol were rinsed out. We pretreated cytoskeletons with 100 μM ATP in PEM plus 20% glycerol for 30 minutes, rinsed out the glycerol and ATP, and then incubated the ATP-pretreated cytoskeletons for 60 minutes in PEM alone. The ATP pretreatment of the Glu MTs in the presence of glycerol had little effect (see above); however, if these Glu MTs were subsequently incubated in PEM without ATP or glycerol, almost all Glu MTs disassembled within 60 minutes (Fig. 7c). Controls incubated in glycerol without ATP and then rinsed and incubated in PEM, contained abundant Glu MTs (Fig. 7d). Thus, ATP hydrolysis can be temporally separated from the disassembly of the Glu MTs, showing that ATP hydrolysis is required only to initiate disassembly. This result suggests that ATP modifies the Glu MT cap, which then permits MTs to depolymerize by subunit exchange.

ATP-treated Glu MTs fail to incorporate tubulin subunits

ATP treatment might also modify the Glu MT cap to allow tubulin subunits to grow off the ends. We tried growing tubulin subunits from ATP-treated Glu MTs (stabilized by glycerol or taxol) but did not observe growth under any conditions. Attempts to induce growth with higher tubulin concentrations, by including ATP during growth, or by incubating for longer, were unsuccessful. Combined treatments (e.g. ATP and salt) also failed to relieve the constraint against growth, although we were limited in the salt concentration we could use, since high salt broke down the MTs. These results raise two possibilities. (1) The Glu MT cap is not removed by ATP treatment, and MTs break down by a mechanism other than plus-end depolymerization (such as severing, or minus-end depolymerization). (2) The cap remains attached to the plusend of the Glu MT, but the ATP treatment alters its activity to allow depolymerization of the MT without permitting subunit incorporation.

The ATP-induced breakdown of Glu MTs is an end-mediated process

We directly viewed ATP-induced Glu MT breakdown to determine the mechanism of ATP disassembly of Glu MTs. Cells with R-tubulin-labeled MTs were extracted and incubated in buffer for 15 minutes to depolymerize Tyr MTs (see Materials and Methods). The remaining stable MTs were imaged by low light-level timelapse recording. We observed fluorescent, stable MTs in extracted cells for at least 30 exposures before detectable photodamage. Introduction of 200 μM ATP caused most MTs to shorten almost immediately (Fig. 8a). The shortening occurred without evidence of fragmentation (Fig. 8a). We observed a number of examples in which shortening MTs straightened out during the course of depolymerization, as if shortening of the MT relieved a constraint on it (see Fig. 8a). This last result in particular shows that the ATP-induced depolymerization of MTs is an end-mediated process. Without addition of ATP, almost all MTs remained intact throughout the recording and did not change length (Fig. 8b). This confirms that the MTs we imaged in these experiments were stable MTs and that shortening was dependent upon added ATP.

Fig. 8.

ATP-induced end-dependent depolymerization of R-MTs in extracted cells. TC-7 cells with microinjected R-tubulin were detergent-extracted, and then incubated in buffer to remove dynamic Tyr MTs. The remaining stable MTs were timelapse recorded. Extracted cells were incubated with PEM containing 200 μM ATP (a, 0-8 minutes) or PEM alone (b, 0-8 minutes). Arrows in the a series show the successive positions of three MT ends as they depolymerize. Incubation time in minutes is indicated in the upper right corner. Bar, 10 μm.

Fig. 8.

ATP-induced end-dependent depolymerization of R-MTs in extracted cells. TC-7 cells with microinjected R-tubulin were detergent-extracted, and then incubated in buffer to remove dynamic Tyr MTs. The remaining stable MTs were timelapse recorded. Extracted cells were incubated with PEM containing 200 μM ATP (a, 0-8 minutes) or PEM alone (b, 0-8 minutes). Arrows in the a series show the successive positions of three MT ends as they depolymerize. Incubation time in minutes is indicated in the upper right corner. Bar, 10 μm.

The rate of ATP-induced depolymerization of Glu MTs determined from our recordings was 2.2±1.4 μm/minute (n=11). For comparison, the rate of depolymerization of dynamic MTs in intact cells was 28.2±13.9 μm/minute (n=13). Other studies report rates of MT depolymerization in vivo of 10-20 μm/minute (Shelden and Wadsworth, 1993; Mikhailov and Gundersen, 1998). We did not measure the rate of depolymerization of Tyr MTs in extracted cells, but since most Tyr MTs disappeared within minutes of extraction, this rate must be >10 μm/minute. Therefore, ATP treatment stimulates Glu MT depolymerization, but does not convert Glu MTs into MTs that depolymerize rapidly like dynamic (Tyr) MTs in vivo or in extracted cells.

ATP-treated Glu MTs lose subunits from their plus ends

The above experiments did not identify the end from which depolymerization of Glu MTs occurred. Two scenarios were possible: (1) subunits were lost from the plus end or (2) subunits were lost from the minus end and the MT was reeled in toward the centrosome. To distinguish between these possibilities, we made a mark on R-labeled Glu MTs in extracted cells by photobleaching a zone of fluorescence before perfusing ATP (see Materials and Methods). If subunits were lost from the plus end, the bleached zone should remain at a fixed point as the MT depolymerizes. If subunits were lost from the minus end, the bleached zone should move in toward the cell center as the MT depolymerizes. We photobleached MTs in 18 cells and detected 42 photobleached MTs with clearly observable ends. In every case when an end shortened, the bleached zone did not move. In 28 of the 42 bleached MTs, it was clear from the orientation of the MT (free end in the cell periphery; other end near the centrosome), that the plus end shortened; in the remaining cases, it was not clear which end was shortening. In at least 18 of the 28 cases (64%) in which plus ends depolymerized, there was a clear stimulation of depolymerization by ATP. (In some experiments, ATP did not stimulate depolymerization of all the MTs during the short time course of the experiment.)

A typical example of ATP-induced depolymerization from an MT end is shown in Fig. 9. Before ATP perfusion, the MT maintains its length (panels ‘0’ and ‘30’), whereas after ATP perfusion, the MT shortens dramatically from its distal end. During shortening, the bleached zone does not move (the bleached zone is most clearly in focus in panels ‘280’ and ‘340’). Fig. 9 also shows that the ATP-stimulated depolymerization continued through the bleached zone (panels ‘420’ to ‘510’), demonstrating that the laser exposure did not damage the MT. In control cells (no ATP), photobleached R-MTs remained stable and did not depolymerize during the course of the experiment.

Fig. 9.

ATP-treated Glu MTs depolymerize from their plus ends. TC-7 cells microinjected with R-tubulin, were detergent-extracted and incubated in PEM to remove dynamic Tyr MTs. Then, a photobleached zone on the remaining stable MTs was created by laser exposure and 200 μM ATP was perfused. Time is indicated in seconds in the upper right hand corner of each panel. An area of the cell with two MTs with clear ends is shown (cell center is on the right). Panel ‘0’ is before photobleaching. A photobleached zone was made on the two MTs at 20 seconds (the photobleached zone can be seen in the ‘20’ second panel by the direct fluorescence emitted during the photobleaching and in the subsequent panels by a zone without fluorescence). After ATP perfusion (at ‘190’), the MT on the top shortens without movement of the bleached zone. The shortening continues through the bleached zone (panels ‘420’ and ‘510’). The lower MT, which has both ends free, shortens through the bleached zone (panel ‘190’) and then continues shortening until it disappears at ‘420’. The black cross hair in the middle of the images obscures some of the fluorescence from the top MT and should not be confused with the photobleached zone. The white vertical line in the images is a digital defect. Bar, 20 μm.

Fig. 9.

ATP-treated Glu MTs depolymerize from their plus ends. TC-7 cells microinjected with R-tubulin, were detergent-extracted and incubated in PEM to remove dynamic Tyr MTs. Then, a photobleached zone on the remaining stable MTs was created by laser exposure and 200 μM ATP was perfused. Time is indicated in seconds in the upper right hand corner of each panel. An area of the cell with two MTs with clear ends is shown (cell center is on the right). Panel ‘0’ is before photobleaching. A photobleached zone was made on the two MTs at 20 seconds (the photobleached zone can be seen in the ‘20’ second panel by the direct fluorescence emitted during the photobleaching and in the subsequent panels by a zone without fluorescence). After ATP perfusion (at ‘190’), the MT on the top shortens without movement of the bleached zone. The shortening continues through the bleached zone (panels ‘420’ and ‘510’). The lower MT, which has both ends free, shortens through the bleached zone (panel ‘190’) and then continues shortening until it disappears at ‘420’. The black cross hair in the middle of the images obscures some of the fluorescence from the top MT and should not be confused with the photobleached zone. The white vertical line in the images is a digital defect. Bar, 20 μm.

Characterization of the ATPase capping activity of Glu MTs

To further characterize the Glu MT capping activity, we screened pharmacological agents to identify those that inhibited the ATP-induced breakdown of Glu MTs. We tested inhibitors of kinases, phosphatases, dynein, kinesins and proteases, all at or above concentrations reported to be effective in inhibiting their specific targets. Only AMP-PNP and vanadate inhibited the ATP-induced breakdown of Glu MTs (see Table 3). AMP-PNP (at 1 mM) almost completely prevented the breakdown of Glu MTs by 100 μM ATP, whereas vanadate (at 1 mM) reduced, but did not completely prevent breakdown (Table 3).

Table 3.

Inhibition of ATP-induced breakdown of Glu MTs by pharmacological reagents

Inhibition of ATP-induced breakdown of Glu MTs by pharmacological reagents
Inhibition of ATP-induced breakdown of Glu MTs by pharmacological reagents

Given our previous results, which implicated protein phosphorylation in the destabilization of Glu MTs (Gurland and Gundersen, 1993), we were surprised that kinase inhibitors did not prevent the ATP-induced breakdown of Glu MTs. To ensure that the inhibitors blocked kinase activity, we tested whether staurosporine, the most broadly effective inhibitor we used, inhibited protein phosphorylation in the extracted cells. We incubated extracted cells with [γ-32P]ATP, collected soluble (released) and cytoskeletal fractions, and analyzed the phosphorylation patterns by SDS-PAGE and autoradiography (see Materials and Methods). Phosphorylation of a limited number of polypeptides was detected in the soluble and cytoskeletal fractions (Fig. 10, lanes S2, C2) and this was completely inhibited by 50 μM staurosporine (Fig. 10, lanes S3, C3). Since 50 μM staurosporine did not inhibit the ATP-induced breakdown of Glu MTs (Table 3), but completely inhibited protein phosphorylation, kinase activity is not required for the ATP-induced disassembly of Glu MTs. Conversely, when AMP-PNP was added during the ATP incubation, no detectable change in the number of phosphorylated polypeptides, or in the degree of phosphorylation, was noted (Fig. 10, lanes S1, C1), despite the fact that AMP-PNP blocked Glu MT breakdown (Table 3).

Fig. 10.

Inhibition of protein phosphorylation in extracted cells by staurosporine, but not by AMP-PNP. Extracted TC-7 cells were incubated with [γ-32P]ATP for 1 hour in the absence (S2, C2) or presence of 1 mM AMP-PNP (S1, C1), or 50 μM staurosporine (S3, C3). Soluble fractions (S1-3) and cytoskeletal fractions (C1-3) were isolated, subjected to SDS-PAGE and autoradiographed. Molecular mass markers are shown on left.

Fig. 10.

Inhibition of protein phosphorylation in extracted cells by staurosporine, but not by AMP-PNP. Extracted TC-7 cells were incubated with [γ-32P]ATP for 1 hour in the absence (S2, C2) or presence of 1 mM AMP-PNP (S1, C1), or 50 μM staurosporine (S3, C3). Soluble fractions (S1-3) and cytoskeletal fractions (C1-3) were isolated, subjected to SDS-PAGE and autoradiographed. Molecular mass markers are shown on left.

The properties of the ATPase regulating Glu MT stability resemble those of kinesin; both prefer ATP, but can use other nucleotide triphosphates, and both are inhibited by AMP-PNP and vanadate (McIntosh and Porter, 1989). We used antibodies against two major kinesins, conventional kinesin (SUK4, H2), and kif 3A (K2.4), but did not observe kinesin on Glu MT ends, although we did detect kinesin staining of vesicular structures in cells (Fig. 11). Neither kinesin was detected in material released from Glu MTs by ATP, even though both were detected in initial extracted cells (not shown). These results suggest that Glu MT capping activity is not due to conventional kinesin or kif3A and instead may be due to a novel kinesin.

Fig. 11.

Kinesin is not detected on the ends of Glu MTs by immunofluorescence. Unextracted TC-7 cells were stained for Glu tubulin (a) and conventional kinesin (SUK 4 antibody; b). The boxed areas in a and b are magnified and overlaid in (c). Bars, 10 μm.

Fig. 11.

Kinesin is not detected on the ends of Glu MTs by immunofluorescence. Unextracted TC-7 cells were stained for Glu tubulin (a) and conventional kinesin (SUK 4 antibody; b). The boxed areas in a and b are magnified and overlaid in (c). Bars, 10 μm.

Relationship between MT stability, capping and detyrosination

This paper provides evidence for a novel MT capping mechanism that functions on interphase MTs and accounts for the selective stabilization of Glu MTs in cells Since the discovery of subsets of MTs with enhanced stability toward treatments that depolymerize MTs (Brinkley and Cartwright, 1975; Thompson et al., 1984) and the correspondence of these MTs with long-lived MTs (Webster et al., 1987a; Schulze et al., 1987), the mechanism responsible for the stabilization has remained enigmatic. The discovery that these MTs contain elevated levels of post-translationally detyrosinated tubulin (Gundersen et al., 1984) suggested that the modification was responsible for the stability. However, now it is clear that the modification is a consequence of MT stability, rather than a cause. Increased detyrosination of tubulin is not sufficient to enhance MT stability in vitro (Skoufias and Wilson, 1998), in extracted cytoskeletons (Khawaja et al., 1987) or in vivo (Webster et al., 1990). Elevated detyrosination is also not necessary for stable MT formation, since stabilized MTs are detectable before increases in detyrosination (Cook et al., 1998). Also, some cell lines do not generate MTs with elevated detyrosination and yet have stable MTs (Gundersen and Bulinski, 1986; Webster et al., 1987b). The capping mechanism we have described does not involve detyrosination as part of the mechanism to stabilize MTs. Instead, it suggests a model in which the MTs become capped first and detyrosinated after stabilization.

Capping of individual MTs can also explain the selective nature of MT stabilization observed in most cells. MTs that encounter active capping factors could be stabilized even though MTs in the vicinity remain dynamic. Localized regulation of such a capping activity must occur to generate the polarized arrays of stable MTs in polarized and differentiated cells (see below). We have recently identified a Rho dependent pathway that stimulates the Rho effector mDia to selectively stabilize MTs (Cook et al., 1998; A. F. Palazzo, T. A. Cook, A. S. Alberts and G. G. Gundersen, unpublished).

Characteristics of the Glu MT cap

Our studies of the Glu MT cap suggest that it exhibits a number of unusual properties and may be composed of a novel protein or protein complex. We showed that the cap is tightly bound to the ends of Glu MTs, since it resists detergent extraction, mild salt and dilution in buffer. Most importantly, we found that the activity of the cap could be modified by ATP hydrolysis, so that it no longer stabilized MTs against dilution-induced disassembly. Our timelapse and photobleaching studies showing that ATP triggered loss of tubulin from the plus end are critical to our interpretation that the cap itself is affected by ATP. Nevertheless, we think that ATP only modifies the cap rather than removes it, since the ATP-treated MTs depolymerized slowly and would not grow when tubulin subunits were added.

It is unclear how ATP modifies the cap. Results with kinase inhibitors and [γ-32P]ATP strongly suggest that a kinase is not involved, and results with nonhydrolyzable analogues argue that energy is required. The glycerol experiments show that the energy of ATP hydrolysis is not required for loss of each tubulin subunit. This leads us to think that ATP hydrolysis induces a conformational change in the cap such that the MT can lose subunits but is still resistant to addition of monomeric tubulin.

It will be important to identify the molecular components of the Glu MT cap in future studies. Localization studies suggest that known MT-end binding proteins are not involved in the Glu MT cap (Table 1). EB1, APC, vinculin focal adhesions and p150Glued are not detected on the ends of most Glu MTs. Other studies have shown CLIP-170 to be specifically localized on dynamic MTs (Perez et al., 1999), so we do not think it is a likely candidate. CLIP-170 is also colocalized with p150Glued, which is not found on a high percentage of Glu MT ends (Vaughan et al., 1999).

Based upon the nucleotide specificity, inhibitor profile and location of the cap on the plus end, we think members of the kinesin superfamily are leading candidates for the cap. Indeed, the behavior of the ATP-treated Glu MT cap is similar to that of kinesin-coated beads attached to MTs ends (Lombillo et al., 1995). Lombillo et al. showed that kinesin beads bound to the plus ends of MTs move with depolymerizing MTs. Importantly, ATP was not required to keep the kinesin beads attached as the MT depolymerized. Similarly, the Glu MT cap remained with the depolymerizing MT, and continued depolymerization did not require ATP.

We tested two members of the kinesin superfamily, conventional kinesin (kif5) and a subunit of type II kinesin (kif3A), and found that neither of them was localized at the ends of Glu MTs nor was solubilized from Glu MTs by ATP treatment. Thus, if a kinesin is involved in capping Glu MTs, it is probably another member of the kinesin superfamily. At least one kinesin, XKCM1, is known to preferentially interact with MT ends (Desai et al., 1999). However, XKCM1 may not be a good candidate for the Glu MT cap, since its principal activity is to induce MT catastrophe. In preliminary experiments, we have not observed localization of MCAK, a mammalian orthologue of XKCM1 (Wordeman and Mitchison, 1995), on the ends of Glu MTs. It will be interesting to test additional family members as good probes for them become available.

Regulation of Glu MT capping

Our finding that micromolar concentrations of ATP stimulate end-wise depolymerization of Glu MTs in extracted cells presents something of a paradox. The ATP concentration in cells is ≥1 mM and yet cellular Glu MTs remain remarkably stable, persisting for up to 16 hours in TC-7 cells with no significant change in length (Webster et al., 1987a). Even at the slow rate of depolymerization we measured for ATP-treated Glu MTs in extracted cells (2.2 μm/minute), an average 50 μm MT would depolymerize in approximately 25 minutes in vivo. We think that ATP triggers Glu MT shrinkage in extracted cells because key regulatory elements that prevent Glu MT breakdown in vivo are removed by detergent extraction. In support of this is the lack of effect of phosphatase and kinase inhibitors. In vivo phosphatase inhibitors result in the rapid depletion of Glu MTs (Gurland and Gundersen, 1993; Merrick et al., 1997) and some kinase inhibitors induce the formation of Glu MTs in vivo (T. A. Cook and G. G. Gundersen, unpublished observations).

Comparison of the Glu MT cap to other MT ‘caps’

Two well-known cases where MTs are capped are kinetochores in the mitotic spindle (Mitchison, 1988) and at the distal ends of axonemes (Miller et al., 1990; Wang et al., 1994). MT capping of a sort may also occur during cell division in yeast, where cytoplasmic MTs are captured at the bud cortex and move the associated nucleus toward the bud neck by depolymerization (Adames and Cooper, 2000). Although captured yeast MTs are much less stable than interphase Glu MTs, the loss of subunits from the captured MTs while they shrink is similar to the shrinking of capped Glu MTs treated with ATP. Kinetochore and axoneme caps contribute to the stability of the attached MTs, as with the Glu MTs cap. Both types of capped MTs are capable of growth and shrinkage at certain times, while remaining associated with their caps (Miller et al., 1990; Mitchison and Salmon, 1992; Wang et al., 1994). These comparisons suggest that capping of MTs may be a general way to couple the energy of MT polymerization/depolymerization in order to move associated cellular structures.

The proteins that are responsible for capping kinetochore and axonemal MTs have not been identified, although motor proteins of both the dynein and kinesin families have been implicated (Yen et al., 1992, Pfarr et al., 1990; Steuer, 1990; Wordeman and Mitchison, 1995). Proteins involved in the yeast MT capture have been identified genetically (Heil-Chapdelaine et al., 1999), yet the molecules responsible for capping the MTs have not been unequivocally identified. The yeast orthologue of EB1, Bim1/Yeb1, has been implicated in MT capture by virtue of its association with MT ends and its interaction Kar9, which is localized at the bud tip (Korinek et al., 2000; Lee et al., 2000). However, it is not yet clear that Bim1/Yeb1 is on the captured MT. That EB1 is not localized on Glu MT ends suggests that, for capped MTs in interphase mammalian cells, EB1 need not be present. A yeast kinesin, kip3, has also been implicated in the capture of yeast MTs (DeZwann et al., 1997; Lee et al., 1999) but there is no known mammalian equivalent. It will be interesting to compare the proteins identified in these various MT caps with the Glu MT cap to determine whether MT capping involves a conserved set of proteins.

The authors thank Dr Guojuan Liao for comments on the manuscript, Dr Alexei Mikhailov for advice on recording fluorescent MTs, and Dr Paul Kronebusch for help with lasing. Dr E. Hilal was instrumental in developing the MT growth assay. A. Infante was supported in part by an NIGMS training grant to the Integrated Program in Cellular, Molecular and Biophysical Studies. This study was supported by NIH grant (GM-42026) and ACS grant (RPG-89-006-13) to G.G.G.

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