Microtubule reconfiguration during axogenesis - Semantic Scholar

Journal of Neurocytology 30, 861–875 (2001)

Microtubule reconfiguration during axogenesis W E N Q I A N Y U 1 , C H A N G Y I N G L I N G 2 a n d P E T E R W. B A A S 1,∗ 1

Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA; 2 Department of Anatomy, University of Wisconsin Medical School, Madison, WI 53706 [email protected] Received 22 April 2002; revised 29 May 2002; accepted 30 May 2002

Abstract When cultured on polylysine, rat sympathetic neurons extend modest lamellae which contain a mass of relatively short nonaligned microtubules. Microtubules display movements, but these movements do not result in any obvious alterations in the overall configuration of the array. Application of a mixture of growth factors called matrigel results in a rapid expansion of the lamellae followed by the outgrowth of axons. Microtubules undergo rapid behavioral changes that result in dramatic alterations in the microtubule array. Microtubules become significantly longer, and extend to the periphery of the lamellae where they invade newly-forming axons. The microtubules align with one another and relative to the cell cortex, and draw together into bundles. Microtubules within a bundle move apart as well, particularly at the tips of developing axons. These observations demonstrate a complexity of microtubule behaviors, some of which can be explained by interactions with actin and/or by forces generated by molecular motor proteins.

Introduction The axon is an elongated cellular process extended from the neuron for the purpose of transmitting information over potentially long distances in the body. The genesis of the axon represents the initial polarization of the neuron, and is a seminal event in neuronal differentiation. The axon is a cylindrical tube of cytoplasm surrounded by cell membrane and tipped by a highly motile structure called the growth cone. Axons can branch extensively during development, and undergo dramatic bouts of retraction as well as elongation as they achieve their final patterns of innervation. The genesis and remodeling of the axon depend on finely orchestrated changes in the cytoskeletal elements that underlie its morphology. Actin filaments are essential for the motility of the growth cone, but elaboration of the axon itself requires the establishment of a highly organized array of microtubules. Microtubules are essential for both the architecture of the axon and for directing organelle traffic along its length. Axogenesis is preceded by the formation of motile lamellae, which give rise to axons as microtubules invade their peripheral regions. The lamellae collapse, and their remnants become growth cones at the tips of the developing axons. During axogenesis, scattered microtubules funnel into the elongated processes and become aligned with one another, thus forming an organized paraxial bundle (Yu & Baas, 1994). What are the mechanisms by which ∗ To

whom correspondence should be addressed.

0300–4864

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2002 Kluwer Academic Publishers

a scattered array of microtubules gives rise to a tightly organized paraxial array during axogenesis? Several studies have shown that microtubules are highly dynamic in newly-forming axons (Ahmad et al., 1993; Kabir et al., 2001), and that these dynamics are important for events such as growth cone navigation (Tanaka & Kirschner, 1995; Challacombe et al., 1997). However, the dynamic properties of the microtubules cannot fully explain the various patterns of organization that characterize different regions of the neuron at different stages of its development. Studies on other cell types have established that microtubules are configured by interactions with other cytoskeletal elements such as actin, by motor proteins that generate forces upon them, and by a variety of other accessory proteins with various mechanisms of action (see for example Sharp et al., 2000; Reilein et al., 2001; Vallee et al., 2001). It appears that the same is true of the neuronal microtubule arrays (Baas, 1999), but the precise behaviors and relevant molecules that configure neuronal microtubules remain poorly understood. In the current study, we have utilized a culture system in which axogenesis occurs very rapidly from broad flat lamellae that are extremely advantageous for both live-cell imaging of microtubules and for electron microscopic analyses. Our immediate goal was to characterize the system and document microtubule behaviors underlying

862 axogenesis so that future functional hypotheses can be tested. Methods CELL CULTURE

Superior cervical ganglia were dissected from newborn rat pups, treated with trypsin and collagenase, and triturated into a single-cell dispersion as previously described (Ahmad et al., 1998). The cells were then plated onto the glass-bottomed well of a 35-mm petri dish into which a 1-cm had been drilled and an acid-washed glass coverslip had been adhered. Prior to plating the cells, the glass-bottomed well had been treated for 3 hours with 0.1 mg/ml polylysine, and then rinsed extensively with water. The cells were plated in a modified L15based medium that maintains pH in normal air (Ahmad et al., 1998). The neurons were then permitted to attach to the substratum and extend lamellae for at least 2 hours and sometimes overnight. Virtually no axonal outgrowth was observed under these conditions over this period of time. Rapid expansion of the lamellae and axogenesis were induced by the addition of matrigel (Collaborative Biomedical Products) diluted 1 : 200–1 : 400 from the stock provided by the company (Slaughter et al., 1997). PREPARATION AND MICROINJECTION OF FLUORESCENT TUBULIN

Tubulin was purified from bovine brain by several cycles of polymerization-depolymerization, and stored as a pellet of microtubules frozen at − 80◦ C. The fluorescent dye 5-(+6)-carboxytetramethyl-rhodamine succinimidyl ester (TMR) (Molecular Probes, Eugene, OR) was coupled to tubulin as previously described (Keating et al., 1997). This labeling procedure resulted in polymerization-competent TMR-tubulin with dye-to-protein ratios of 0.8–1.2. TMRtubulin was diluted to 4 mg/ml in injection buffer (100 mM PIPES, 0.5 mM MgCl2 , pH 6.9), aliquoted, and stored in liquid nitrogen. Before injection, aliquots were thawed and centrifuged at 21,000 g (15,000 rpm) at 4◦ C for 5 minutes (Eppendorf Model 5402, Hamburg, Germany) to remove tubulin aggregates. TMR-tubulin was microinjected into neurons using the Eppendorf Microinjector 5242/Manipulator 5170 as described in Dent et al. (1999). LIVE - CELL IMAGING

Imaging of the microtubule array within living neurons was performed essentially as previously described (Dent et al., 1999). The imaging system included an Axiovert 135 M inverted microscope (Zeiss Incorporated, Thornwood, NY) equipped with the a long bandpass rhodamine filter set, a Keller port to maximize the amount of emitted light to the camera, electronically-controlled shutters (Uniblitz, Vincent Associates, Rochester, NY), and a slow-scan liquid-cooled charge-coupled device (CCD) (Photometrics PXL, Tucson, AZ) equipped with a Kodak KAF-1400 chip. Illumination during all epifluorescence imaging was reduced to 10–25% of the output of the light source by placement of neutral density filters (Chroma Technology, Brattleboro, VT) in the light path. Cells were maintained at 36◦ C with an airstream incubator (Nicholson Precision Instruments, Bethesda, MD). Neurons

Y U , L I N G and B A A S that had been injected with fluorescent tubulin were imaged with a 100× /1.3 NA Fluar objective (Zeiss). Images were acquired every 10–20 sec, with 100–1000 msec exposures, under the low light level conditions described above. Fine focus was manually controlled with an LEP MAC 2000 focus controller (Ludl Electronic Products, Hawthorne, NY). The CCD camera and shutters were controlled by Metamorph 2.5 software (Universal Imaging, West Chester, PA) running on a Pentiumbased computer (Datastor, Boulder, CO). Images were collected at a 500–800 kHz transfer rate, as described in detail in Dent et al. (1999). All images were saved to the hard drive in 12-bit format.

FLUORESCENCE MICROSCOPY ON FIXED NEURONS

The cytoskeletal arrays of the neurons were also visualized with fluorescent probes applied after fixation. For this, cultures were extracted and fixed simultaneously for 15 minutes in a solution containing 4% paraformaldehyde, 0.15% glutaraldehyde, and 2% Triton X-100 in a buffer termed PHEM (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, 2 mM MgCl2 , pH 6.9). Cells were rinsed extensively in PBS, and then exposed for one hour at room temperature to a solution containing a 1 : 100 dilution of Cy3-conjuaged anti-β-tubulin antibody (Sigma) together with a 1 : 10 dilution of Alexa Fluor 488 phalloidin (Molecular Probes, Eugene, OR). After extensive rinsing in PBS, cultures were mounted in an anti-fade medium, and then visualized and photographed with the microscope system described above (without the neutral density filters).

ELECTRON MICROSCOPY

Transmission electron microscopy was used to generate higher-resolution images of the configuration of microtubules relative to other microtubules and relative to actin filaments. Conventional preparative procedures for transmission electron microscopy are notoriously poor at preserving actin filaments, and in our experience, can often result in images of rather poor contrast in studies on cultured sympathetic neurons. In the present study, we have used a protocol that optimizes for actin preservation, and also results in images of much better contrast than procedures used in most of our previous studies. This protocol was developing using modifications from other protocols reported to help preserve actin filaments and enhance contrast (for example, Tilney et al., 1996). After removing the medium, cultures were fixed for 2 minutes at 37◦ C in a solution containing 2% glutaraldehyde and 2 mg/ml tannic acid, after which the cultures were placed on ice for an additional 28 minutes of fixation. After three rinses for 5 minutes each in 0.2 M cacodylate (pH 6.8), cultures were postfixed with 1% OsO4 in 0.1 M cacodylate for 20 minutes, rinsed twice for 2 minutes each in 3.6% NaCl, rinsed twice for 2 minutes each in water, and then stained for 30 minutes with 5% uranyl acetate in water. Cultures were then dehydrated in progressively increasing concentrations of ethanol, and embedded in Durcupan. Thin sections were obtained with an Ultracut-E Ultramicrotome (ReichertJung, Germany). Sections were 0.1 µm thick, which resulted in almost the complete thickness of the lamellae in one section. Sample were visualized and photographed with a JEOL CX100 electron microscope (JEOL, Peabody, MA).

Microtubule reconfiguration during axogenesis Results FEATURES OF MATRIGEL - INDUCED AXOGENESIS

The overall goal of this study was to document changes in microtubule configuration during axogenesis, and to do so in a system that permits high-resolution microscopic analyses that will prove useful for future mechanistic studies. In order to accomplish this, it was necessary to use a neuronal cell culture system that would permit us to discern microtubules with great clarity, and one in which axogenesis occurs rapidly. We found that a culture regime initially described by Slaughter and colleagues (1997) is well-suited for these studies. These authors found that the addition of matrigel to rat sympathetic neurons that had been plated on polylyine induced an extremely rapid and robust outgrowth of multiple axons. We observed an additional advantage of the system for our studies on the earliest phases of axogenesis; namely that axons elongated from extraordinarily broad and flat lamellae. These lamellae are typically less than 0.15 µm in thickness in their more peripheral regions, and hence circumvent classic problems with imaging neuronal microtubules created by the thickness of the sample (see Baas, 2000). Neuronal cell bodies and typical axons are not flat, and are densely packed with microtubules, thus preventing analyses that require a great deal of spatial resolution by light microscopy. Electron microscopy is better suited for thicker samples, but requires serial reconstruction of consecutive thin sections in order to appreciate the

863 complexity of the microtubule array (see for example Yu & Baas, 1994; Yu et al., 1994). The thin flat lamellae generated by the matrigel regime provide a unique opportunity to study microtubule behaviors in living neurons, and also to visualize most or all of the microtubules in a single thin section with the electron microscope. We modified the procedure of Slaughter and colleagues in that we used an L15-based medium that would maintain pH in normal air (see Ahmad et al., 1998). Figure 1 shows phase-contrast images of rat sympathetic neurons two hours after plating, and various times after the subsequent addition of matrigel. By two hours on polylysine, the cell bodies have already extended modest lamellae. These lamellae are quite flat, even prior to adding the matrigel (Fig. 1A). Within 30 minutes of adding the matrigel, the lamellae begin to expand (Fig. 1B), often encompassing 3–4 times more surface area than prior to adding the matrigel (Fig. 1C). Still within the first hour of adding the matrigel, thin cylindrical axons tipped by flat growth cones can be viewed extending from the periphery of the lamellae. Axonal growth ensues rapidly, and the lamellae associated with the cell body gradually collapse as the axons continue to develop (Fig. 1D–H). This same general pattern of axogenesis has also been observed when sympathetic plated on polylysine are exposed to laminin (Rivas et al., 1992; Tang & Goldberg, 2000). The similarity observed under these different regimes suggests to us that axogenesis in these neurons occurs by a

Fig. 1. Phase-contrast images of rat sympathetic neurons two hours after plating, and after addition of matrigel. By two hours on polylysine, the cell bodies have extended modest lamellae (panel A). By thirty minutes after adding the matrigel, the lamellae begin to expand (panel B), often encompassing 3–4 times more surface area than prior to adding the matrigel (panel C). Still within the first hour of adding the matrigel, thin cylindrical axons tipped by flat growth cones extend from the periphery of the lamellae. The lamellae associated with the cell body gradually collapse as axons develop (panels D–H). Bar, 30 µm.

864 stereotyped sequence of events. In addition, and very importantly for our studies, the matrigel response is faster, more dramatic, more consistent from cell to cell in the culture, and produces broader flatter lamellae than laminin or any other axonal growth regime that has been reported. Shown in Figure 2 are higher magnification phasecontrast images of lamellae in the early phases of axogeneseis. In our previous experience observing the flattened growth cones and branch points of cortical neurons, we learned that the darker regions in phasecontrast micrographs usually represent areas of high microtubule density and alignment (unpublished observations, see also Dent et al., 1999). In Figure 2A, a darkened thin filopodium can be seen (marked by an arrow) forming at the periphery of the lamella; this presumably represents a region into which aligned microtubules are beginning to invade. Figure 2B shows early phases of the lamella crimping and dividing into multiple nascent axons. Figure 2C shows a dark swirl around the edge of the lamella (marked by an arrow). A similar dark swirl gives rise to a nascent axon in Figure 2D (marked by an arrow). Figure 2E shows more crimping of the lamellae, and Figure 2F shows how such crimping gives rise to multiple axons. Figure 2G shows a less common phenomenon with a very dark band (marked by an arrow) extending through the lamella (rather than

Y U , L I N G and B A A S hugging its edge as is more common) and leading into a nascent axon. Figure 2H shows an early axon with a large growth cone extending from a lamella while the lamella itself is also crimping in the region directly continuous with the cell body. Assuming that the darkened regions indeed correspond to areas dense with aligned microtubules, these images provide initial clues as to how microtubules change their distribution and configuration during the formation of axons. LIVE - CELL IMAGING OF MICROTUBULE BEHAVIORS

In order to directly appreciate the organization of microtubules in these neurons, we microinjected them with fluorescently-labeled tubulin, which rapidly incorporates into the microtubules. Unlike immunostains for tubulin, this approach also permits us to observe the changes in the microtubule array that occur over time. Imaging began one hour after microinjection to permit sufficient time for the fluorescent tubulin to incorporate into microtubule polymers. Because the lamellae are so flat, it is possible to image the microtubules with far more clarity than would be possible with typical cultured neurons in which the resolution is obstructed by both the thickness of the microtubule array as well as a great deal of haze caused by the free fluorescent tubulin. Resolution was similar or better than that

Fig. 2. Higher magnification phase-contrast images of lamellae in the early phases of axogeneseis. In panel A, a darkened thin filopodium can be seen (marked by an arrow) forming at the periphery of a lamella. Panel B shows crimping and dividing of a lamella into multiple nascent axons. Panel C shows a dark swirl around the edge of the lamella (marked by an arrow). A similar dark swirl gives rise to a nascent axon in panel D (marked by arrow). Panel E shows more crimping of a lamella. Panel F shows how such crimping gives rise to multiple axons. Panel G shows a less common phenomenon with a very dark band (marked by arrow) extending through the lamella and leading into a nascent axon. Panel H shows an early axon with a large growth cone extending from a lamella while the lamella itself is also crimping in the region directly continuous with the cell body. Bar, 17 µm.

Microtubule reconfiguration during axogenesis obtained in our previous study on the flattened growth cones and branch points of cultured cortical neurons (Dent et al., 1999). Fluorescence generated by the small amount of free tubulin within the flattened lamellae does not obstruct good resolution of the polymers, but fortuitously delineates the edges of the lamellae. We observed 88 neurons before matrigel-treatment and at various times after the addition of matrigel. The results were remarkably consistent from cell to cell at similar points in the regime. Figure 3 shows images obtained over a period of 90 minutes of a neuron cultured on polylysine with no addition of matrigel. The microtubules in the lamella appear as a dense mass of mostly non-aligned polymers, some of which (marked by the

865 larger arrow) swirl around parallel to but not close to the edge of the lamella (rather than invading into it as occurs after matrigel treatment, see later). At the 12 minute time point, a bud of the lamella expands outward, but the microtubules do not move into the bud, which subsequently collapses back, as can be seen in the 57 minute time point. The dense microtubule array appears to splay and open-up in one area at 57 minutes, but even by 90 minutes, it is apparent that this does not result in any major reconfiguration of microtubules, which still do not align or invade to the edge of the lamella. Thus any movements that may occur are ultimately futile in terms of generating notable changes in microtubule organization. Major alterations in the microtubule array occur after the addition of matrigel, as evidenced by Figure 4, which shows a typical matrigel-induced reconfiguration of microtubules. Most of the microtubules are now aligned with their distal ends directed toward the edge of the lamella. At 0 minutes, two particularly dense bundles of microtubules are apparent at the outer edges of the lamella, which has already begun to crimp into axons. Importantly, these bundles of microtubules are not surrounded on all sides by cell membrane, indicating that microtubule bundling cannot be explained simply by the membrane collapsing around unbundled microtubules. (Similar observations were reported of neurons treated with laminin; see Tang & Goldberg, 2000). These microtubule bundles that run along the edges of the lamellae are common at this stage of axogenesis (observed in almost all of the samples we monitored), and are reminiscent of the phase-dark areas showed in Figure 2C and D. By 20 minutes, one of these microtubule bundles has given rise to an early axon, with the lamella crimping around it to form a cylinder. The other bundle shows slower development over 20 and 40 minutes, but has produced a second distinct axon by 60 minutes. The region of the lamella

Fig. 3. Live-cell images obtained over a period of 90 minutes of a neuron cultured on polylysine with no addition of matrigel. Microtubules in the lamella appear as a dense mass of mostly non-aligned polymers, many of which swirl around parallel to but not close to the edge of the lamella. At 12 minutes, a bud of the lamella expands outward (marked by an arrow), but the microtubules do not move into the bud, which subsequently collapses back, as can be seen in the 57 minute time point. The dense microtubule array appears to splay and open-up in one area at 57 minutes, but even by 90 minutes, it is apparent that this does not result in any major reconfiguration of microtubules, which still do not align or invade to the edge of the lamella. Sometimes aster-like formations of microtubules could be observed (see lower right area in the lower two panels). In the 90 minute panel, the larger arrow points to the swirl of microtubules that appear to be swept back from the leading edge of the lamella, while the smaller arrow points to an aster-like formation of microtubules. Bar, 10 µm.

866 between the two early axons has still not collapsed into a discrete axon by 60 minutes, which is reflected by the fact that the microtubules are generally aligned but not yet tightly bundled. In the growth cones at the tips of

Y U , L I N G and B A A S the newly-forming axons, the microtubules once again splay apart from the bundle (as seen of the second axon to form in the 60 minute time point). Figure 5 provides more insight into the bundling of microtubules. In this sequence, we see a region of the lamella that has started to crimp to form an axon; there are several aligned microtubules in this region at 0 minutes, but they have not yet formed a tight bundle. At 7, 18, and 30 minutes, the microtubules can be seen gradually moving together into a bundle as this region of the lamella is transformed into a cylindrical axon tipped by a growth cone. Figure 6 provides more insight into the splaying of microtubules from a bundle in a newly-formed growth cone. A bundle of microtubules extends into a large growth cone and bends around as seen at 0 minutes. At 18 minutes, the bundle can be seen to loosen, and individual microtubules begin to splay off the bundle into the body of the growth cone. At 29 minutes, more splaying can be observed, with a portion of the bundle breaking free from the main bundle. At 44 and 46 minutes, additional microtubule movements during the splaying can be observed. The movements can be quite rapid, as evidenced by marked differences observed at these latter time points, which are only 2 minutes apart. Images obtained by fixing cultures and then staining with a fluoresently-labeled tubulin antibody were similar in appearance to the live-cell images of neurons injected with fluorescent tubulin. For this reason, we did not focus a great deal of attention on fixed cell images. However, an additional point can be gleaned from images in which we double-labeled for actin filaments. Figure 7A shows a newly-formed microtubule bundle in green, surrounded by the cortical actin in red. The bundling of microtubules (such as that shown in Fig. 5) has occurred prior to the cortical actin completely compressing around the edges of the nascent axon. The axon

Fig. 4. Alterations in the microtubule array after the addition of matrigel. Most of the microtubules are aligned with their distal ends directed toward the edge of the lamella. At 0 minutes, two particularly dense bundles of microtubules are apparent at the outer edges of the lamella, which has already begun to crimp into axons. These bundles are not surrounded on all sides by cell membrane. By 20 minutes, one of these microtubule bundles has given rise to an early axon, with the lamella crimping around it to form a cylinder. The other bundle shows slower development over 20 and 40 minutes, but has produced a second distinct axon by 60 minutes. The region of the lamella between the two early axons has still not collapsed into a discrete axon by 60 minutes; the microtubules are generally aligned but not yet tightly bundled. In the growth cones at the tips of the newly-forming axons, the microtubules once again splay apart from the bundle (as seen of the second axon to form in the 60 minute time point). Bar, 15 µm.

Microtubule reconfiguration during axogenesis

Fig. 5. Microtubule bundling. In this sequence, a bud of a lamella has started to crimp to form an axon. There are several aligned microtubules at 0 minutes, but they have not yet formed a tight bundle. At 7, 18, and 30 minutes, the microtubules can be seen gradually moving together into a bundle as this bud of a lamella is transformed into a cylindrical axon tipped by a growth cone. Bar, 12 µm.

shown in Figure 7 displays a distal region in which the cortical actin has constricted around the microtubule bundle, and a more proximal region where it has not. Another observation gleaned from examination of hundreds of fixed cell images was the frequent appearance (both before and after matrigel addition) of aster-like formations within dense arrays of microtubules (see Fig. 7B). These asters were also sometimes observed in the live-cell images (in Fig. 3, note the aster toward the right side of the lamella at the 57 and 90 minute time

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Fig. 6. Splaying of microtubules from a bundle in a newlyformed growth cone. A bundle of microtubules extends into a large growth cone and bends around as seen at 0 minutes. At 18 minutes, the bundle loosens, and individual microtubules begin to splay off the bundle into the growth cone. At 29 minutes, more splaying is observed, with a portion of the bundle breaking free from the main bundle. At 44 and 46 minutes, additional microtubule movements during the splaying can be observed. The fact that movements can be very fast is evidenced by marked differences observed between these latter time points, which are only 2 minutes apart. Bar, 12 µm.

points; marked by the smaller arrow in the 90 minute panel). Similar aster-like formations are commonly observed in vitro when motors and microtubules are combined (Compton, 2001).

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Fig. 7. Fluorescence labeling of cultures after fixation. Microtubules are shown in green and actin filaments are shown in red. Panel A shows a newly-formed bundle of microtubules similar to that shown in Figure 5. The contrast of the actin-staining (in red) has been digitally increased so that the perimeter of the lamella is as clear as possible. Notice that the actin-rich cortex has not yet compressed tightly around the bundle in the more proximal region of the axon. Panel B shows an aster formation of microtubules, marked by an arrow. Bar (both panels), 5 µm. ELECTRON MICROSCOPIC ANALYSES

To obtain higher resolution images of microtubules and to better appreciate their configuration relative to one another and to actin filaments, we performed transmission electron microscopy. We analyzed sections of roughly 50 cells before matrigel treatment, 30 minutes after, and 1 hour after the addition of matrigel. As with the live-cell imaging, the results were very consistent from cell to cell at similar stages in the regime. As noted in the Materials and Methods section, we used a preparative procedure that optimizes for the preservation of actin filaments and provides better contrast of the microtubules than routine procedures used in our previous studies on sympathetic neurons. Even so, it was our strong impression that actin filaments preserved well when they were part of dense bundles but preserved poorly when they were not (as evidenced by the paucity of detectable actin filaments other than those in bundles). There were dramatic changes in ultrastructure upon matrigel treatment, both with regard

Y U , L I N G and B A A S to microtubules and the actin bundles. In cells not treat with matrigel, most of the actin bundles swirled around the periphery of the lamella, parallel to its edge (see Fig. 8A). Sometimes small regions at the edge of the lamella extended outward as a small “bud’’ (as can be seen at the top of the 12 minute panel in Fig. 3; marked by an arrow). Within these buds, the actin bundles were oriented parallel to the long axis of the bud. In the case of both types of actin orientation (which were essentially perpendicular to one another), microtubules were generally sparse in the actin-rich regions, but they were not entirely excluded. Some microtubules were aligned parallel to the actin bundles (see larger arrow in Fig. 8B), while other microtubules ran more perpendicular to the actin bundles (see two smaller arrows in Fig. 8B). In deeper regions of the lamellae (farther from the periphery), actin filaments were not detected, and the microtubules were generally short and scrambled in appearance (Fig. 8C and D). The ultrastructure of cells exposed to matrigel was markedly different. There was a substantial diminution of actin filament levels, and a dramatic reorganization of the filaments that persisted. Instead of a dense and continuous array of cortical actin as observed before matrigel treatment, at 30 minutes after adding matrigel, there were discrete rib-like bundles of actin filaments that extended into the filopodia that extended from the lamellae (Fig. 9A–C). These bundles sometimes ended blindly deeper in the lamella, but often curved around the periphery of the lamellae from one filopodium to the next (as in Fig. 9C; see arrow). Microtubules were often seen aligning with the path of the actin bundles (as in Fig. 9D and E; see arrows), and as the filopodia expanded into nascent axons, microtubules began to occupy the space. In addition, the microtubules took on the general alignment of the actin bundles that previously dominated these regions (as in Fig. 9F, see arrow). With additional time in matrigel, at one and two hour time points, the actin bundles became restricted to the growth cones at the tips of developing axons. The microtubules underwent a variety of dramatic changes with increasing time in matrigel. By one hour, the microtubule profiles were generally many times longer than in cells not treated with matrigel (see all panels in Figs. 10 and 11). Given that almost the entire thickness of the peripheral regions of a lamella could be captured within a single thin section, we can conclude that this difference in microtubule length is not an artifact of sectioning. Another very consistent observation is that many of the microtubules took on a cortical distribution that mirrors the earlier distribution of actin filaments. Figure 10A shows microtubules swirling in a tight bundle around the edge of the lamella one hour after matrigel treatment, in a very similar distribution to that of the actin filaments after 30 minutes shown in Figure 9C. In Figure 10A and also Figure 10B, it is apparent that those microtubules that do not tightly


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Fig. 8. Electron microscopic analyses on neurons not treated with matrigel. Panel A shows that most of the actin bundles swirl around the periphery of the lamella, parallel to its edge. In buds of the lamella that extend outward, the actin bundles are oriented parallel to the direction of outgrowth of the bud (panel B). Microtubules are generally sparse in the actin-rich regions, but are not entirely excluded. Some microtubules align parallel to the actin bundles (an example is marked by the larger arrow in panel B), while other microtubules are more perpendicular to the actin bundles (an example is marked by the two smaller arrows in panel B). In deeper regions of the lamellae farther from the periphery, actin filaments are not detectable, and the microtubules are generally short and scrambled in appearance (panels C and D.) Bar, 0.75 µm.

hug the edge of the lamella are scattered and less organized relative to one another. In Figure 10B, the transition from organized to less-organized microtubules is less distinct than in Figure 10A, probably illustrating the recruitment of scattered microtubules into the bundle. Figure 10C shows that the swirl of microtubules around the edge of the lamella continues directly into a newly-formed axon. Microtubules are also recruited into the axon from deeper regions of the lamella (see lower part of Fig. 10C, where microtubules not hugging the periphery also enter the axon). Figure 11A and B show deeper regions of the lamella, not near its edge or near regions of axonal formation. Even in these regions, and throughout all regions of the lamellae, the microtubules are longer and more organized relative to one another than in cells not treated with matrigel. It is very common for small groups of microtubules to be aligned with one another, and notably, the microtubules can be seen “drawing’’ together into small bundles. That is, the microtubules are often farther apart at one end of the bundle than the other (see in particular, Fig. 11B, arrowheads). This suggests that bundle formation is the result

of a zippering process. Figure 11C shows a microtubule bundle around the edge of a lamella that expands into a primitive growth cone. The microtubules can be seen to splay apart from the bundle in what appears to be a reverse-zippering process. Such splaying was common within growth cones. Discussion The present studies demonstrate that microtubules undergo rapid behavioral changes during axogenesis. Prior to axogenesis, the microtubule array undergoes movements, but these movements are essentially futile in that they do not result in any major alterations in the organization or distribution of microtubules. Although a small number of individual microtubules can enter and even align with the actin bundles at the periphery of the lamellae, the vast majority of the microtubule mass does not penetrate the dense cortical actin array. Small buds of the lamellae occasionally extend outward, and when this happens, actin bundles align parallel to the direction of outgrowth rather than

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Fig. 9. Electron microscopic analyses of cells exposed to matrigel for 30 minutes. A substantial diminution of actin levels and a dramatic reorganization of the filaments that persist are apparent. Discrete rib-like bundles of actin filaments extend into the filopodia (panels A–C). These bundles sometimes end blindly deeper in the lamella, but often curve around the periphery of the lamella from one filopodium to the next (panel C; see arrow). Microtubules often align with the path of the actin bundles (as in panels D and E; see arrows). As the filopodia expand into nascent axons, microtubules begin to occupy the space. In addition, the microtubules take on the general alignment of the actin bundles that previously dominated these regions (as in panel F, see arrow). Bar, 0.75 µm.

circumferentially around the edge of the lamella. However, there is no detectable increase in the levels of microtubules that invade into the bud, and the bud rapidly collapses back. The microtubules do not align or bundle with one another in any obvious way, except where they appear to be swept back in unison as they

approach the actin-rich periphery. Robust alterations in both the microtubule and actin arrays are apparent within 30 minutes of axogenesis. The dense actin array gradually diminishes, giving way to smaller realigned bundles. The microtubules then show dramatic realignment with one another, particularly in regions where


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Fig. 10. Electron microscopic analyses showing further changes in microtubule configuration one hour after addition of matrigel. The microtubule profiles are many times longer than in cells not treated with matrigel (see also Fig. 11 and compare with Fig. 8). Many of the microtubules show a cortical distribution that mirrors the earlier distribution of actin filaments. Panel A shows microtubules swirling in a tight bundle around the edge of the lamella, in a very similar distribution to that of the actin filaments after 30 minutes shown in Figure 9C. In panels A and B, it is apparent that those microtubules that do not tightly hug the edge of the lamella are scattered and less organized relative to one another. In panel B, the transition from organized to less-organized microtubules is less distinct than in panel A, probably illustrating the recruitment of scattered microtubules into the bundle. Panel C shows the swirl of microtubules around the edge of the lamella continuing directly into a newly-formed axon. Microtubules are also recruited into the axon from deeper regions of the lamella (see lower part of panel C). Bar, 1.0 µm.

the actin had previously realigned, namely filopodia and the periphery of the lamellae. Bundling of microtubules is apparent prior to the actin cortex or the cell membrane constricting around them, indicating a

true microtubule–microtubule interaction. Particularly large bundles of microtubules form at the periphery of the lamellae, and funnel into newly forming axons. The microtubules within these bundles spread apart once

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Fig. 11. Electron microscopic analyses showing additional features of microtubule configuration after one hour in matrigel. Panels A and B show deeper regions of the lamella, not near its edge or near regions of axonal formation. The microtubules are longer and more organized relative to one another than in cells not treated with matrigel. Small groups of microtubules are aligned with one another, and the microtubules can be seen “drawing’’ together in small bundles in a zipper-like fashion. A particularly good example is marked by arrowheads in panel B. Panel C shows a microtubule bundle around the edge of a lamella that expands into a primitive growth cone (at the right side of the panel, see arrowheads). The microtubules can be seen splaying apart from the bundle. Bar, 0.75 µm.

again within the growth cones at the tips of the axons. We refer to these microtubule behaviors as alignment, zippering, and splaying. These behaviors, which we have also observed in the growth cones of more highly developed axons (Dent et al., 1999), do not occur by wholesale depolymerization and repolymerization. Live-cell imaging shows gradual transitions from scattered to bundled microtubules as well as from bundled to splayed arrays; in no case did we ever observe widespread loss of microtubules followed by the appearance of new microtubules with markedly different configurations. Electron micrographs show scattered microtubules coming together like a zipper, rather than polymers aligned

along their entire length as one might expect of copolymerizing microtubules. Thus we would conclude that many of the microtubule behaviors that occur during axogenesis are not readily explained by dynamics, but are at least in part due to bona fide movements. In an earlier study on matrigel-induced axogenesis in sympathetic neurons, Slaughter and colleagues (1997) documented the transport of microtubules from the cell body into newly growing axons. They showed that this type of microtubule movement is both ongoing and robust. In the present study we did not seek to revisit the issue of whether individual microtubules are transported into the axon, but rather, we sought to understand how combinations of movements could produce major

Microtubule reconfiguration during axogenesis organizational changes during the earliest phases of axogenesis. Our studies document alignment, zippering, and splaying as key factors in the transformation of a scattered microtubule array into a highly ordered one during axogenesis. Studies in vitro, on various cell types, and especially on mitotic cells indicate that forces generated by motor proteins can configure microtubules into various kinds of arrays (for review and discussion see Sharp et al., 2000). For example, it has been shown that purified motors added to microtubules can organize them into both asters and aligned bundles (Nedelec et al., 1997; Surrey et al., 2001). Our frequent detection of asters in neuronal lamellae may or may not have functional significance, and the possibility remains that these asters might merely be the result of random crossings of microtubules. On the other hand, given that aster formation is a direct result of motor interactions with microtubules, the asters we observed might be an indication that motors are indeed generating forces on the microtubules within these arrays. Zippering is also known to be a common result of motor-driven forces on microtubules. Cytoplasmic dynein or minus-enddirected kinesins can zipper microtubules toward their minus-ends either by interacting directly with neighboring microtubules or by moving in conjunction with an accessory protein that associates with the motor and with neighboring microtubules (Merdes et al., 2000; Compton, 2001). Plus-end-directed motors can zipper microtubules toward their plus ends, as has been directly observed with the kinesin termed Eg5, which sorts microtubules with regard to their polarity and then zippers them into bundles (Walczak et al., 1998). It is unknown whether neurons express minus-enddirected kinesins with zippering properties, but we have shown that Eg5 is highly expressed in developing neurons and tends to concentrate on microtubules at the leading edge of regions of new growth (Ferhat et al., 1998). Splaying of microtubules could occur by local inactivation of the motor that zippers, or by a more active process whereby microtubules are drawn away from one another by yet another motor-driven force. The idea of microtubule bundling by motor proteins is very different from the classic view that microtubules are bundled by non-motor fibrous proteins that engage neighboring polymers along their length. However, the motor-based hypothesis is consistent with the long-standing dissenting view that the fibrous microtubule-associated proteins are not actually the culprits that draw neuronal microtubules together into bundles (for discussion see Weisshaar et al., 1992). Indeed, microtubule bundles generated by overexpression of fibrous microtubule-associated proteins such as tau have a paracrystalline appearance that is quite different from that of microtubule bundles in neurons expressing these proteins at physiological levels (see for example Baas et al., 1991; Chen

873 et al., 1992). However, at their physiological level, the fibrous microtubule-associated proteins might bundle microtubules more appropriately. In favor of a role for fibrous microtubule-associated proteins, Teng and colleagues (2001) have reported defects in microtubule bundling in mice with impaired expression of one or more of the fibrous microtubule-associated proteins. It may be that motor proteins are needed to generate microtubule bundles via zippering, and that non-motor microtubule-associated proteins are needed to stabilize the interactions between microtubules once the bundle has formed, and to determine the spacing between microtubules within the bundle. Our studies also suggest a potential role for actin filaments in organizing microtubules. Prior to axogenesis, the dense array of actin at the periphery of the lamellae seems to act as a barrier to the invasion of the microtubule mass. This may be partially due to the configuration of the actin filaments (mostly perpendicular to the path of future axonal outgrowth), and partially due to a retrograde flow of actin by myosin motors, as has been observed in pausing growth cones (Lin et al., 1996) and fibroblasts (Waterman-Storer & Salmon, 1997). Indeed, as noted above, the microtubules appear to be swept backward in unison from the leading edge. An early event in axogenesis that precedes the redistribution of microtubules is a global reorganization of the actin meshwork at the periphery of the lamellae. Actin bundles reorient paraxially within buds that give rise to new axons, but are much less dense than the bundles within the transient buds that sometimes arise prior to axogenesis. In addition, the actin bundles hug the periphery of the lamellae, and sometimes are continuous with the paraxial bundles leading out into the nascent axons. These actin bundles have essentially the same orientation and location that the microtubules will subsequently assume, which is consistent with the idea that actin bundles may act as tracts for the transport of microtubules by cytoplasmic dynein (Pfister, 1999; Ahmad et al., 1998). Interestingly, though, it was not common to observe microtubules in direct side-byside alignment with actin bundles, as one would expect if the microtubules were pushing directly against the actin. We suspect that there are alterations in the actin array during axogenesis that are simply not visible in our micrographs. There appears to be a great deal of actin debundling in regions of the cytoplasm that become microtubule-rich, and this may render the individual filaments more difficult to preserve and/or detect. At present, we cautiously interpret the redistribution of microtubules along previously established actin patterns as evidence that actin may be a substrate for microtubule transport, but a great deal of mystery still surrounds this issue. While our studies and commentary have focused on microtubule movements and interactions with other microtubules and actin, it is important to stress that

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microtubule dynamics are also critically important. There is abundant evidence for the importance of microtubule assembly, microtubule stabilization, microtubule destabilization, and the influence of a variety of regulatory proteins. For example, it has been shown that microtubules must be highly dynamic in the region of the growth cone in order for axons to navigate appropriately in response to environmental cues (see for example Challacombe et al., 1997). Microtubule destabilizing proteins such as stathmin and SCG10 help maintain a portion of the microtubules in their dynamic form in the axon (Pellier-Monnin et al., 2001), which is also laden with proteins such as STOP, MAP1b, doublecortin, and tau, which have putative microtubulestabilizing properties (Baas et al., 1994; Guillaud et al., 1998; Mack et al., 2000; Caspi et al., 2000). Stabilization of domains on the microtubules is important to ensure the preservation of their plus-end distal polarity pattern (Baas & Black, 1990; Baas & Ahmad, 1992). In addition, microtubules undergo changes in length as the axon develops (Yu & Baas, 1994; Yu et al., 1994), as evidenced here by their marked elongation during the earliest phases of axogenesis. Thus we would conclude that microtubules undergo dramatic movement-based behaviors during axogenesis, but that these movements occur against a backdrop of other critically important behaviors. One of the most interesting features of our data is the speed by which axogenesis occurs upon addition of matrigel. This suggests that the proteins needed for axogenesis are already present, and are modified rapidly to produce the requisite alterations in cytoskeletal organization. It has already been established that biochemical cascades involving phosphorylation target microtubule-associated proteins such as MAP1b during axonal development (Gordon-Weeks & Fischer, 2000), and it seems reasonable that other relevant proteins (such as force-generating motors) might also be key targets for these cascades. Future studies will be aimed at using this culture regime to address specific hypotheses on the molecules and mechanisms that underlie the microtubule behaviors reported here.

Acknowledgments This work was supported by two grants to Peter W. Baas from the National Institutes of Health. We thank Daniel Buster, Carley Sauter, and Yan He (of our laboratory) for assistance and helpful discussions.

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