Experimental Neurology 189 (2004) 112 – 121 www.elsevier.com/locate/yexnr
Myosin II activity is required for severing-induced axon retraction in vitro Gianluca Gallo * Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA Received 8 December 2003; revised 28 April 2004; accepted 14 May 2004 Available online 1 July 2004
Abstract Understanding the mechanistic basis of the response of neurons to injury is directly relevant to the development of effective therapeutic approaches aimed at the amelioration of nervous system damage. Axons retract in response to severing. We investigated the mechanism of axon retraction in response to severing in vitro, testing the hypothesis that actomyosin contractility drives severing-induced axon retraction. Axon retraction commenced within 5 min following severing and correlated with actin filament accumulation at the site of severing. Depolymerization of actin filaments prevented retraction, demonstrating that actin filaments are required for severing-induced axon retraction. Direct inhibition of myosin II, using blebbistatin, minimized axon retraction in response to severing. Blocking RhoA-kinase (ROCK), a modulator of myosin II activity, inhibited axon retraction. Similarly, inhibiting myosin light chain kinase (MLCK) with a cellpermeable pseudo-substrate peptide also inhibited axon retraction. These data demonstrate that myosin II activity is required for severinginduced axon retraction in vitro, and suggest myosin II as a target for therapeutic interventions aimed at minimizing retraction following severing in vivo. D 2004 Elsevier Inc. All rights reserved. Keywords: Actin; RhoA-kinase; Myosin light chain kinase; Axotomy; Cytoskeleton; Injury
Introduction Injury to the nervous system results in retraction of axons (Finger and Almli, 1985; Houle and Tessler, 2003). During the period of axon retraction, the site of injury develops into an inhibitory territory for axon extension (Gimenez y Ribotta et al., 2002). Thus, if axons could be blocked from undergoing the initial injury-induced retraction, then the axon tips would be at a point of advantage when regenerative attempts are made. The distance axons would be required to regenerate would be minimized, and axons may start regenerating before the development of myelinderived inhibitory signals. Therefore, promoting prompt regeneration and minimizing the time window during which neurons attempt to reconnect to their targets are of relevance in abating the long-term effects of injury. Although the retraction of inappropriately targeted axon branches is a fundamental mechanism of neurodevelopment and has detrimental effects following injury, the mechanism * Department of Neurobiology and Anatomy, Drexel University College of Medicine, 2900 Queen Lane, Philadelphia, PA 19129. Fax: +1-215-843-9082. E-mail address:
[email protected]. 0014-4886/$ - see front matter D 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.expneurol.2004.05.019
of axon retraction is not well understood (Raff et al., 2002). Recent investigations indicate that it is an active process mediated by alterations in motor protein-based force generation. Ahmad et al. (2000) report that blocking the activity of the microtubule-associated motor protein dynein results in actomyosin-dependent axon retraction. Similarly, axon retraction in response to the pharmacological stabilization of actin filaments, or treatment with Ephrin-A2, is dependent on myosin II-driven contractility (Gallo et al., 2002). Myosin II is regulated by myosin light chain kinase (MLCK) and RhoA-kinase (ROCK). MLCK directly phosphorylates myosin regulatory light chains and activates myosin motor activity. ROCK activity, driven by the GTPase RhoA, increases phosphorylation of myosin regulatory light chains by phosphorylating and inhibiting the myosin light chain phosphatase, and possibly directly phosphorylating the myosin regulatory light chains (Luo, 2000). The introduction of constitutively active RhoA in neurons results in axon retraction and extracellular ligand– receptor-based signals that cause axon retraction do so in a RhoA- and myosin IIdependent manner (e.g., Gallo et al., 2002; Jalink et al., 1994; Wylie and Chantler, 2003). The regulation of RhoA activity has also been shown to underlie axon retraction in the developing Drosophila nervous system (Billuart
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et al., 2001). The mechanism of axon retraction in response to severing has not been elucidated, and it is not known whether it shares similarities with forms of axon retraction induced by extracellular signals. We tested the hypothesis that severing-induced axon retraction requires myosin II activity. This investigation demonstrates that axotomy-induced axon retraction requires myosin II-driven contractility. These data suggest myosin-II and its regulatory kinases as targets for therapeutic approaches aimed at preventing axon retraction following direct physical injury.
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In vitro axon severing
Embryonic day (E) 10 chicken embryos were used as a source of dorsal root ganglia (DRG), E7 embryos were used to produce retinal explants. Embryos were dissected, and DRGs and retinal explants obtained and cultured as previously described (Gallo et al., 2002). Briefly, explants were cultured in 1 ml of defined-medium (F12H with additives; Invitrogen Inc., Carlsbad CA). DRG explants were supplemented with 20 ng/ml nerve growth factor (R&D Systems, Minneapolis, MN). Explants were cultured in video dishes manufactured by drilling a 14-mm diameter hole in the bottom of 60-mm tissue culture dishes and affixing glass coverslips to cover the hole using aquarium sealant (Ernst et al., 2000). The glass surface of video dishes was coated overnight with 25 Ag/ml laminin (Invitrogen) overnight at 39jC before use in culturing. Similarly, for some experiments, substrata were coated 100 with Ag/ml of poly-Dlysine (70 KD MW; Sigma) in borate buffer and washed three times with PBS before use in culturing. Retinal and DRG explants were cultured for 1 and 2 days, respectively, before use in axon-severing experiments to allow axons to extend to lengths between 1 and 2 mm.
For severing experiments, explants were cultured in video-dishes and placed on the microscope stage for 20 min before use in experiments. The culture was then scanned to identify isolated axons. Only axons that exhibited a ‘healthy’ uniform caliber axon were used in single-axon-severing experiments. Severing of axons was performed 700 –1000 and 1500 –2000 Am from the point of origin of the axons extending out from the retinal and DRG explants, respectively. For severing, borosilicate glass capillaries (AM Systems, Carlsborg WA; catalog # 625500) were pulled to fine tips using a Pul-1 micropipette puller (World Precision Instruments, Sarasota FL). Tipped glass capillaries were sealed at the end that was not pulled using aquarium sealant to prevent back fill of the capillary when placed in the medium. Before use in an experiment, the pulled capillary tips were dipped in a solution of 2 mg/ml bovine serum albumin (Sigma) in PBS to minimize the adhesion of cell surfaces to the glass during severing. Capillary needles were mounted on a mechanical micromanipulator (Leica Inc., Bannockburn, IL). The tip was then brought into contact with the substratum and axons severed by maintaining the position of the tip steady while moving the stage relative to the tip. The stage was moved manually in a continuous swift motion resulting in uniform severing of axons. Cases where the axon was displaced laterally by the capillary tip during severing were excluded. The distance retracted by axons following severing was measured using computer-assisted image-analysis by determining the distance between the position of the distal tip of the proximal segment of the axon at the time of severing and the position of the severed tip after a period of time. Thus, the absolute distance the tip retracted in Am/unit time was measured. A minimum of 15 axons was monitored following severing in each experimental group.
Microscopy
Immunocytochemistry
Live imaging experiments were performed on a Zeiss 135M microscope equipped with an AxioCam CCD camera (Zeiss, Gottingen Germany). Live cultures were observed using a 20 phase objective. Time-lapse acquisition was driven by Zeiss AxioVision software. Axons were observed for 20 min following severing in vitro with an image acquired every minute. The stage was heated to 39 – 40jC using an ASI-400 air curtain incubator system (NevTek, Burnsville VA). Quantitative analysis of retraction was performed using the interactive measurement module of the AxioVision software. Imaging of fluorescently labeled preparations was performed on a Zeiss 200 M microscope using a 100 objective. All images and time-lapse sequences were stored digitally. Analysis of the distance axons retracted was performed using Zeiss AxioVision software.
For experiments investigating actin filament distribution vis-a`-vis axon retraction, individual axons were severed, filmed, and fixed at specified times after severing by adding glutaraldehyde directly to the medium at a final concentration of 0.25%. Cultures were fixed for 15 min and subsequently treated with 2 mg/ml sodium borohydride (Sigma, St. Louis, MO) before staining. Actin filaments were stained using rhodamine-phalloidin (Molecular Probes, Eugene OR; 8/100 Al of staining solution) and counterstained with DioC(6)3 (Molecular Probes; 30 s with 2.5 Ag/ml) to reveal axon morphology, as previously described (Ernst et al., 2000). Cultures were then mounted in NoFade medium (Ernst et al., 2000) and stored at 20jC. For determination of the relative distribution of actin filaments and total tubulin in severed axons, cultures were fixed with 0.25% glutaraldehyde and stained with 1:100 FITC-conjugated
Materials and methods Tissue culture
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DM1A anti-tubulin (Sigma) and rhodamine phalloidin in 10% goat serum containing 0.1% Triton X-100 for an hour. Quantitative analysis of actin filament and tubulin staining Images were acquired as dual channel (actin filaments and tubulin) stacks in identical register. Acquisition parameters were set for each channel such that no pixel was at the saturation value. The visible distal-most extent of either of the two staining patterns was used as the starting point for determination of the staining intensity. Total staining intensity was then determined within 5-Am lengths of the axon for both actin filaments and tubulin proceeding in a disto-proximal manner until either the axon overlapped with another axon or the axon being measured reached the border of the screen. Total staining intensity was obtained using background subtraction as previously described (Ernst et al., 2000). Reagents Latrunculin-A was obtained from Molecular Probes. Y27632 was from Calbiochem (San Diego, CA). Blebbistatin was purchased from Toronto Research Chemicals Inc. (North York, Ontario Canada). The cell-permeable myosin light chain kinase (MLCK) inhibitory peptide was designed
based on the MLCK pseudosubstrate sequence reported by Kemp et al. (1987). For the present experiments, the MLKCinhibitory peptide was synthesized in tandem with the cell permeable peptide penetratin (Lindgren et al., 2000) inserted at the amino terminus of the MLCK-peptide (American Peptide Company, Sunnyvale, CA; full sequence, Arg-GlnIle-Lys-Ile-Trp-Phe-Gln-Asn-Arg-Arg-Met-Lys-Trp-LysLys-Lys-Arg-Arg-Trp-Lys-Lys-Asn-Phe-Ile-Ala-Val). The penetratin peptide alone (Qbiogene, Carlsbad CA) was used as a control for experiments using the penetratin-MLCK inhibitory peptide.
Results Response of axons to severing in vitro The behavior of individual axons, growing from retinal or dorsal root ganglion (DRG) explants, was monitored for a 20- to 40-min period following severing. Axons were initially severed 80 –120 Am behind the tip. DRG axons raised in either nerve growth factor or brain-derived neurotrophic factor exhibited similar responses. Retinal and DRG axons exhibited similar responses to severing and are thus discussed together. Axons retracted following severing (Fig. 1A). The distal ends of the proximal axon
Fig. 1. Severing-induced axon retraction. (A) Sequence of images from a time-lapse video of a dorsal root ganglion axon cultured in NGF before severing ( 1 min; minutes are relative to the time of severing) and after severing (0 and 20 min). The black arrow at 20 min denotes the distance over which axon retraction was measured. The white arrowhead shows the distal segment of the severed axon that has contracted into a bead and is not motile. The white arrows at 20 min denote two growth cones that had not been severed and extended during the period the severed axon retracted. (B) Graph showing the absence of a relationship between the distance behind the growth cone at which severing occurred (x-axis) and the distance the proximal segments of severed axons retracted during a 20-min post-severing imaging period ( y-axis).
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started to shorten by 2 –3 min after severing. The distal portion of the axon separated from the cell body by the severing also underwent marked changes. When the axon was severed less than 30– 40 Am from the tip, the whole distal segment contracted into a bulb-like structure during the imaging period (Fig. 1A). The distal parts of axons severed >50 Am from the tip also underwent proximodistal contraction, but growth cones remained motile throughout the 20-min observation period. For the rest of our experiments, we used DRG explants raised in nerve growth factor. We investigated whether the distance behind the tip at which severing occurred influenced the rate of retraction of the proximal segment after severing. Fig. 1B shows a graph of the distance axons retracted following severing as a function of the distance from the tip at which the severing occurred. No relationship between these variables is evident (r2 < 0.001), indicating that within the range tested (10 – 200 Am proximal to the growth cone), the potential for retraction following severing is equal along the length of the axon. The average distance that axons retracted during a 20-min period after severing was 56 F 7 Am (mean F SEM). Ninety percent of axons retracted greater than 5 Am following severing. For subsequent experiments, we adopted a standardized protocol of 20-min imaging following severing at approximately 80 –100 Am proximal to the growth cone.
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filaments are prominent at the growing tip, the growth cone. As revealed by phalloidin staining, in the axon shaft, actin filaments are not only relatively sparse, but also present in the form of bright puncta (Fig. 2A). To determine the organization of actin filaments in severed axons as a function of time following severing, individual axons were
Attachment of the axon shaft to the substratum decreases severing-induced axon retraction To determine whether substratum attachment of the axon shaft affects the distance axons retract following severing, DRG explants were cultured on poly-lysinecoated substrata. Axons growing on poly-lysine attach to the substratum along the majority of their length and with significantly greater strength due to the highly positive charge of poly-lysine (Pollack and Liebig, 1990). Comparison of the distance axons retracted following severing when cultured on laminin versus poly-lysine revealed that on poly-lysine axons retracted 23% of the distance of axons cultured on laminin-coated substrata (13 F 6 Am/ 20 min, mean F SEM, n = 12). On poly-lysine, 66% of axons retracted greater than 5 Am following severing. Therefore, the degree of attachment to the substratum can modulate axon retraction following severing. For the remainder of our experiments, we used laminin-coated substrata because laminin allows axons to retract more vigorously and thus facilitates data acquisition and investigation of the underlying cellular mechanisms. Actin filaments accumulate at the distal tip of the proximal segment of severed axons We investigated the organization of actin filaments in axons before and after severing. In growing axons, actin
Fig. 2. Formation of an F-actin ‘cap’ following axon severing. (A) Example of F-actin organization, determined by phalloidin staining, in an exemplary unsevered axon. Note the relatively diffuse axonal F-actin distribution and puncta of F-actin (arrowheads). (B) For the purpose of direct comparison, this image is representative of the F-actin distribution in the severed axon shown retracting in Fig. 1A. The axon was fixed 20 min after severing. Note the presence of bright F-actin staining at the tip of the proximal segment of the severed axon (white arrow). We refer to the F-actin accumulation at the tips of severed axons as an F-actin ‘cap’. F-actin in the rest of the severed axon is comparable to that in unsevered axons (compare to panel A). The white arrowheads facing one another indicate the position of severing. The gray arrowhead denotes the F-actin distribution in one of the growth cones shown to continue extending in 1A. The white arrowhead denotes the contracted distal axon, as shown in Fig. 1A. (C) Semiquantitative determination of actin ‘cap’ formation following severing. Axons were severed and monitored by time-lapse microscopy before fixation. For these experiments, we consciously sought axons that failed to retract even up 20 min after severing to determine the status of F-actin ‘cap’ formation in axons that did not retract. Thus, the distribution of the percentage of axons that retracted in this data set is biased to obtain data from a variety of axonal responses to severing. For presentation, axons were divided into two groups based on the response to severing, axons that retracted (Ret), and those that did not (No Ret). The presence of an F-actin ‘cap’ was determined blind to the time after severing. Black bars denote the number of axons scored as exhibiting an F-actin ‘cap’, white bars show the number of axons without a detectable F-actin ‘cap’. Note that almost all axons that retracted after an initial 5-min period exhibited ‘caps’ while the majority of axons that failed to retract did not exhibit ‘caps’. Axons that retracted during the first 5 min after severing exhibited the highest relative percentage of retracting axons without an F-actin ‘cap’, suggesting the initial phase of retraction may be Factin independent.
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severed and their response monitored. Axons were fixed at 1 –5, 5 – 10, or 10– 20 min following severing and stained with phalloidin to reveal actin filaments. The organization of actin filaments in axons was then correlated to the time when the axon was fixed following severing and whether the proximal axon segment had initiated retraction. Actin filament accumulation was evident in the distal tip of the majority of proximal axon segments that underwent retraction following severing (Fig. 2B). Conversely, accumulation of actin filaments was minimal or not detectable in axons that had not initiated retraction at the time of fixing (Fig. 2C). The absence of actin filament accumulation was observed even in axons fixed 10 – 20 min following severing that had not started to retract. These observations demonstrate a correlation between the accumulation of actin filaments at the distal tip of the proximal axon segment and the commencement of axon retraction following severing. To determine whether the accumulation of actin filaments at the proximal tips of severed axons may reflect a nonspecific accumulation of proteins in the retracting axon, we severed axons and subsequently stained them with phalloidin to visualize actin filaments and a tubulin antibody. Because we did not extract the membrane at the time of fixation, the tubulin staining pattern reveals the microtubule polymer as well as cytosolic tubulin. In double-stained axons, we observed a clear accumulation of actin filaments distal to accumulation of microtubules or cytosolic tubulin (Figs. 3A –D) in 13/16 axons, as determined by qualitative observation and quantitative analysis of the staining patterns (Fig. 3E). The three axons that exhibited overlap between tubulin and actin filament staining at the tip also exhibited a bulbous tip morphology, indicating a swelling of the membrane (not shown). These observations demonstrate that the accumulation of actin filaments at the tips of severed axons is not attributable to a nonspecific accumulation of the cytoskeleton or of cytosolic proteins.
Fig. 3. Accumulation of F-actin at the tip of severed axons relative to tubulin distribution in severed axons. Panel A shows an example of a severed (blue arrowhead, white arrow denotes direction of retraction) axon tip that underwent retraction stained for actin filaments (red; F-actin) and tubulin (green). Note that there is not significant overlap between the actin filament and tubulin staining. A nonsevered axon with a growth cone is shown for comparison (white arrowhead). Panels B – D show another example of a severed axon tip stained for actin filaments and tubulin. Panels B and C show the actin filament and tubulin staining patterns, respectively. The white arrow in A shows the direction of retraction. Panel D shows the overlay of the two staining patterns. The blue arrow in B and C denotes the distal-most extent of tubulin staining. Note that the actin filament staining is distal to the tubulin staining. (E) A representative example of the quantitative analysis of actin filament and tubulin staining. This example was obtained from the axon shown in B – D. The axon and the regions sampled in 5-Am intervals are shown above the graph and each region is denoted by numbers 1 – 7, with 1 being the most distal portion of the axon. Note that the actin filament staining (red line) peaks distal to the tubulin (green), indicating accumulation ahead of the accumulation of tubulin staining. White bars in A and D represent 5 Am.
Depolymerization of actin filaments inhibits axotomy-induced axon retraction Actomyosin contractility requires myosin II and actin filaments. Thus, actin filaments were pharmacologically depolymerized before axon severing to determine if the accumulation of actin filaments that correlates with axon retraction reflects a requirement for actin filaments during axon retraction. Latrunuclin-A depolymerizes actin filaments by binding
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to actin monomers and rendering them polymerization incompetent (Coue et al., 1987), thus resulting in filament depolymerization due to the endogenous high rate of filament turnover. Additionally, monomers bound by latrunculin-A are unavailable for polymerization following severing. Cultures were treated with 2 AM latrunculin-A for 5 min before axon severing. As previously reported (Gallo et al., 2002), this treatment regime resulted in the depolymerization of >90% of actin filaments detectable by phalloidin staining in growth cones and axons (not shown). Axons severed in the presence of latrunculin-A underwent minimal retraction in response to severing (Fig. 4). These data demonstrate that actin filaments are required for severing-induced axon retraction.
other myosins (e.g., myosin I, V, or X; Straight et al., 2003; Yarrow et al., 2003). First, we determined the dose of blebbistatin that effectively disrupted myosin II-dependent stress fibers in chicken fibroblasts, and found that treatment with 50 AM blebbistatin maximally disrupted myosin IIdependent stress fibers (not shown). In addition, 50 AM blebbistatin completely blocked myosin II-dependent bleb formation from apoptotic chicken kidney cells in vitro (not shown), consistent with the report of Straight et al. (2003). To directly determine if myosin II activity is required for severing-induced axon retraction, DRG cultures were treated with 1–50 AM blebbistatin before axon severing. Blebbistatin inhibited axon retraction following severing in a dose-dependent manner (Fig. 5A), demonstrating that myosin II activity contributes significantly to retraction. Additionally, the distal segment of severed axons underwent minor contraction in the presence of blebbistatin (Fig. 5A). Quantitative determination of the distance axons retracted following severing revealed that 50 AM blebbistatin inhibited severing-induced retraction by 88% (Fig. 5B). At 25 AM blebbistatin decreased axon retraction following severing by 52% (Fig. 5B). Treatment with 1 AM blebbistatin did not alter the response of axons to severing (Fig. 5B). The dose dependence of blebbistatinmediated inhibition of axon retraction following severing is in accordance with previously published concentrations required to inhibit myosin II in biochemical and in vivo assays (Straight et al., 2003) and in stress fiber dissolution bioassays using chicken primary fibroblasts (50% dissolution of stress fibers at 10 AM; not shown).
Inhibition of myosin II blocks severing-induced axon retraction
RhoA-kinase and myosin light chain kinase contribute to severing-induced axon retraction
Blebbistatin is a recently characterized cell-permeable inhibitor of myosin II ATPase activity that does not inhibit
Myosin II is regulated by RhoA-kinase and myosin light chain kinase. The role of these kinases in severing-induced
Fig. 4. Quantitative assessment of the distance axons retracted following severing in the presence of the F-actin depolymerizing drug latrunculin-A. Data are normalized to the distance retracted by control axons (56 F 7 Am/ 20 min). Latrunculin-A decreased the distance axons retracted by 95% ( P < 0.00001; Welch t test one-tailed).
Fig. 5. Direct block of myosin II activity inhibits severing-induced axon retraction. (A) Sequence of images from a time-lapse of two axons treated with 50 AM blebbistatin for 1 h before severing. Note that axon a1 fails to retract and axon a2 undergoes minimal retraction (compare to Fig. 1A, * denotes site of severing). Axon a1 was severed near the growth cone. Similar to control axons, the distal fragment of a1 initially contracted (arrowhead at 10 min, compare to the distal fragment in Fig. 1A). However, unlike the distal segments of control axons severed near the growth cone, during the next 10 min, the blebbistatin-treated distal axon segment elongated in two directions (arrowheads at 20 min) and exhibited active filopodial extension. (B) Quantitative determination of the distance axons retracted following severing in control and blebbistatin-treated axons (1-h pretreatment). *P < 0.01, **P < 0.0001 relative to no blebbistatin treatment, Welch t test one-tailed.
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axon retraction was tested by treating cultures with pharmacological inhibitors before axon severing. Treatment with 20 AM Y-27632, a dose that completely blocks Ephrin-A2induced retraction in chicken retinal neurons (Gallo et al., 2002) and inhibits myosin light chain phosphorylation (Wahl et al., 2000), decreased the distance axons retracted following severing by 50% (Fig. 6A). These observations indicate that Rho-kinase activity contributes to severinginduced axon retraction. We tested the role of MLCK in axon retraction by using a peptide-inhibitor of MLCK (Kemp et al., 1987). This peptide is expected to have significantly greater specificity than pharmacological reagents because it acts as a pseudosubstrate for MLCK and we previously found this peptide to block MLCK in experiments introducing the peptide into neurons by trituration loading (Gallo et al., 2002). The MLCK-inhibitory peptide was rendered cell permeable by the addition of the penetratin sequence at the N-terminus (see Materials and methods). Hereafter, we will refer to the cell-permeable peptide as MLCKpep. Similar to the reported
Fig. 6. Myosin light chain kinase and RhoA kinase contribute to severinginduced axon retraction. (A) Quantitative assessment of retraction following severing under conditions of inhibited Rho-A kinase activity (1h pretreatment with 20 AM Y-27632) or myosin light chain kinase (1h pretreatment with 1 Ag/ml MLCKpep). Y-27632 and MLCKpep inhibited axon retraction by 52% and 59% relative to controls, respectively ( P < 0.001 in both cases). (B) 1 and 5 Ag/ml MLCKpep inhibit axon retraction ( P < 0.0003 and P < 0.0000001, respectively). Combined treatment with Y27632 + 1 Ag/ml MLCKpep further inhibited retraction relative to Y-27632 alone ( P < 0.001) and, to a lesser extent, MLCKpep alone ( P = 0.1). All comparisons using the Welch t test one-tailed.
effects of a different cell permeable MLCK-inhibitory peptide (Schmidt et al., 2002), 5 Ag/ml MLCKpep stopped axon extension and caused a partial simplification of growth cone morphology (not shown). Treatment of cultures with 5 Ag/ml MLCKpep blocked severing-induced retraction (Fig. 5B). MLCKpep (1 Ag/ml), at a concentration that has minimal effects on growth cone morphology (not shown), inhibited retraction to an extent comparable to the inhibition of Rho-kinase (Fig. 5B). Treatment with both 20 AM Y-27632 and 1 Ag/ml MLCKpep inhibited axon retraction to a greater degree than Y-27632 alone, and to a lesser degree to MLCKpep alone (Fig. 5C). These data suggest that both MLCK and ROCK contribute to axon retraction.
Discussion Retraction is an early response of axons to injury (reviewed in Houle and Tessler, 2003). While the phenomenon of axon retraction following severing has been previously described (Bird, 1978; Chuckowree and Vickers, 2003; Hill et al., 2001; Houle and Jin, 2001; McHale et al., 1995; Oudega et al., 1999; Pallini et al., 1988; Raff et al., 2002), the mechanistic basis of this form of retraction is not understood. To elucidate the mechanism of axon retraction following direct physical injury, we tested the hypothesis that this form of retraction is based on actomyosin contractility. Axon retraction following severing correlated with the accumulation of F-actin at the distal end of the retracting axon. Directly blocking myosin II ATPase activity, using blebbistatin, inhibited severing-induced axon retraction. Inhibiting ROCK pharmacologically or MLCK with a cell permeable specific peptide-inhibitor also attenuated retraction. Thus, severing-induced axon retraction in vitro involves a mechanism that requires actomyosin contractility. F-actin accumulates at the tips of the proximal segments of axons following severing. Axons fixed before the commencement of retraction did not exhibit F-actin accumulation, while axons fixed during retraction contained F-actin accumulation. A previous ultrastructural analysis identified a filamentous network at the tips of severed axons that is likely reflective of the F-actin accumulation we observed (Bird, 1978). Is the F-actin accumulation due to local polymerization of filaments or the recruitment of existing filaments? Our data cannot strictly differentiate between these two possible mechanisms. However, axons contain sparse dynamic actin filaments that undergo turnover on a scale of minutes (Gallo et al., 2002). Thus, if actin filaments are being recruited to the tips of severed axons from more proximal or distal regions, then filaments in the severed axon may also be relatively less dynamic than filaments in unsevered axons. It seems more likely that the tip of a severed axon acquires greater potential for F-actin polymerization than the intact axon, thus resulting in the accumu-
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lation of F-actin. An increase in F-actin polymerization at the distal end of a severed axon may reflect an attempt by the axon to form a new growth cone following severing. Growth cones and axon shafts contain myosin II (Bridgman et al., 2001; Brown and Bridgman, 2003; Rochlin et al., 1995). In growth cones, the majority of F-actin is found in the peripheral domain and does not greatly overlap with myosin II except at the transition zone between the peripheral and central domains. Myosin II can regulate growth cone morphology and axon extension (Bridgman et al., 2001; Diefenbach et al., 2002; Wylie et al., 1998), and the rapid turnover of F-actin in growth cones attenuates the ability of myosin II to generate actomyosin-based contractility that would otherwise result in axon retraction (Gallo et al., 2002). Therefore, given the presence of myosin II in axons, the F-actin accumulation in the distal axon following severing overlaps with myosin II and provides a basis for the generation of a contractile force. The in vitro observations suggest it may be possible to decrease axon retraction following injury in vivo using pharmacological reagents that selectively target myosin II or its regulatory kinases, Rhokinase and MLCK. Indeed, effective inhibition of the RhoA pathway to promote axon regeneration has been accomplished in vivo (reviewed in Ellezam et al., 2002). Wahl et al. (2000) demonstrated that inhibition of the RhoAROCK pathway in retinal ganglion cells abolishes myosin light chain phosphorylation. These considerations suggest that it is feasible to inhibit myosin II activity in in vivo model systems. In addition, inhibition of RhoA signaling decreases cell death at the site of injury (Dubreuil et al., 2003), suggesting that inhibition of the RhoA axis may have multiple protective effects following injury. The effects of inhibiting myosin II directly at the site of injury on neurons and glial cells will require future investigation. While our data demonstrate a role for myosin II and its upstream regulatory kinases in axon retraction following severing, this study does not address the issue of whether myosin II activity in the axon is increased by injury. It is possible that events downstream of severing may activate myosin II in axons above endogenous levels. For example, severing has been shown to cause an influx of calcium ions into the axon (Spira et al., 2001) and MLCK is activated by calcium-calmodulin (Pfitzer, 2001). Our data provide evidence that MLCK activity is required for axon retraction following severing, suggesting that calcium influx after injury may result in the activation of MLCK. Consistent with this possibly, severing DRG axons in calcium-free saline prevents axon retraction (data not shown). Thus decreasing calcium levels at the injury site may inhibit axon retraction by lowering MLCK activity. Alternatively, severing may activate the RhoA pathway, resulting in increased ROCK activity. The determination of the activation of these pathways in living severed axons will require the direct visualization of RhoA/ROCK and MLCK activity in axons during sever-
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ing. These experiments will require the use of fluorescence resonance energy transfer (FRET) sensors for RhoA and MLCK in living neurons (Chew et al., 2002; Yoshizaki et al., 2003). Dubreuil et al. (2003) have demonstrated RhoA activation in vivo for up to 7 days in neurons following injury to the spinal cord, suggesting that activation of the RhoA-ROCK-myosin axis during injury may contribute to long-term injury-induced axon retraction. Future work will have to further address the issue of the activation of myosin II and its regulatory kinases following injury. Conversely, we have previously shown that endogenous myosin II activity is sufficient to drive axon retraction in response to the pharmacological stabilization of actin filaments (Gallo et al., 2002). Thus, following severing axons may have an intrinsic ability to undergo retraction in the absence of injury-induced increases in the levels of myosin II activity. Studies of axon retraction following injury in vivo measured the distance of the severed axon tips from the site of injury at 1 week to 2 months following injury (Hill et al., 2001; Houle and Jin, 2001; Oudega et al., 1999; Pallini et al., 1988). Estimating the distance axons retracted over a 20-min period, similar to the observation period used in the present study, during prolonged times in vivo one obtains a rate of approximately 0.6 –1 Am/20 min. This rate is 50-fold slower than that observed in the present study on a laminin substratum. To our knowledge, no data are available on the rate of axon retraction immediately after injury in vivo. Thus, it is possible that the retraction observed in this study reflects an initial rate of retraction that may decrease with time, as described in vivo by Houle and Jin (2001). Our experimental conditions do not allow us to determine long-term rates of retraction, and the embryonic axons used in this study eventually regenerate following severing in vitro (unpublished observations). Alternatively, embryonic axons may exhibit faster retraction rates than developmentally older axons. However, we suggest that the differences in retraction rates in vitro and in vivo reflect differences between the environments of axons. Unlike in vitro, in vivo axons make numerous contacts and are in a three-dimensional matrix of cells and extracellular matrix that may slow the rate of retraction by providing strong adhesive contacts along the length of the axon. In support of this consideration, we found that axon retraction on the highly adhesive substratum poly-lysine occurred at 23% the rate of retraction on laminin. It will be of interest to determine the time course of axon retraction following injury in vivo on a time scale of hours after severing. Additionally, investigations of the ultrastructure and cytoskeletal organization of axons retracting in vivo should assist in further delineating the mechanistic basis of axon retraction in response to injury. It is also possible that the in vitro observations on rapid retraction following injury may reflect an initial retraction of the axon that proceeds through a different mechanism than prolonged axonal
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retraction. Determination of mechanism of axon retraction in vivo will require taking into account numerous factors not included in in vitro model systems such as the relationships between axons and glial cells and the presence of a complex extracellular matrix. Thus, numerous factors not taken into account in vitro may alter the extent and mechanism of axon retraction in vivo. In conclusion, this study identifies myosin II and its regulatory kinases as potential targets for minimizing axon retraction following injury in vivo. RhoA-kinase activity has been demonstrated to be required for the block of axon extension by myelin-derived inhibitory cues (reviewed by Ellezam et al., 2002). To our knowledge, it is not known whether blocking MLCK also attenuates the inhibitory effects of myelin on axon extension. However, MLCK is upregulated in regenerating axons (Jian et al., 1996), and several reports indicate that MLCK is required for axon extension (Jian et al., 1994; Ruchhoeft and Harris, 1997; Schmidt et al., 2002). Therefore, inhibition of MLCK may not be an ideal target for blocking retraction as it could also hinder regenerative attempts. Conversely, inhibiting RhoAkinase potentiates axon extension improving regeneration (Ellezam et al., 2002; Fournier et al., 2003) and survival of cells at the injury site (Dubreuil et al., 2003). In light of the present findings, these considerations indicate that inhibiting the regulation of actomyosin activity through RhoAkinase in injured axons may also promote regenerative attempts by minimizing the initial retraction and facilitating subsequent attempts at re-extension.
Acknowledgments The author thanks Dr. P.C. Letourneau (University of Minnesota) for providing laboratory space during the initial phase of this research, Dr. R. Loudon and Dr. M. Murray (Drexel College of Medicine) for critical reading of the manuscript, and Ms. L. Silver (Drexel College of Medicine) for excellent technical assistance. This work was supported by grants to G.G. from the Spinal Cord Research Foundation/Paralyzed Veterans of America (#2199) and the National Institute of Health (NS043251-01).
References Ahmad, F.J., Hughey, J., Wittmann, T., Hyman, A., Greaser, M., Baas, P.W., 2000. Motor proteins regulate force interactions between microtubules and microfilaments in the axon. Nat. Cell Biol. 2, 276 – 280. Billuart, P., Winter, C.G., Maresh, A., Zhao, X., Luo, L., 2001. Regulating axon branch stability: the role of p190 RhoGAP in repressing a retraction signaling pathway. Cell 107 (2), 195 – 207. Bird, M.M., 1978. Microsurgical transection of small nerve fibre bundles in vitro. Cell Tissue Res. 190, 525 – 538. Bridgman, P.C., Dave, S., Asnes, C.F., Tullio, A.N., Adelstein, R.S., 2001. Myosin IIB is required for growth cone motility. J. Neurosci. 21, 6159 – 6169.
Brown, J., Bridgman, P.C., 2003. Role of myosin II in axon outgrowth. J. Histochem. Cytochem. 51, 421 – 428. Chew, T.L., Wolf, W.A., Gallagher, P.J., Matsumura, F., Chisholm, R.L., 2002. A fluorescent resonant energy transfer-based biosensor reveals transient and regional myosin light chain kinase activation in lamella and cleavage furrows. J. Cell Biol. 156, 543 – 553. Chuckowree, J.A., Vickers, J.C., 2003. Cytoskeletal and morphological alterations underlying axonal sprouting after localized transection of cortical neuron axons in vitro. J. Neurosci. 23, 3715 – 3725. Coue, M., Brenner, S.L., Spector, I., Korn, E.D., 1987. Inhibition of actin polymerization by latrunculin A. FEBS Lett. 213, 316 – 318. Diefenbach, T.J., Latham, V.M., Yimlamai, D., Liu, C.A., Herman, I.M., Jay, D.G., 2002. Myosin 1c and myosin IIB serve opposing roles in lamellipodial dynamics of the neuronal growth cone. J. Cell Biol. 158, 1207 – 1217. Dubreuil, C.I., Winton, M.J., McKerracher, L., 2003. Rho activation patterns after spinal cord injury and the role of activated Rho in apoptosis in the central nervous system. J. Cell Biol. 162, 233 – 243. Ellezam, B., Dubreuil, C., Winton, M., Loy, L., Dergham, P., Selles-Navarro, I., McKerracher, L., 2002. Inactivation of intracellular Rho to stimulate axon growth and regeneration. Prog. Brain Res. 137, 371 – 380. Ernst, A.F., Gallo, G., Letourneau, P., McLoon, S.C., 2000. Stabilization of growing retinal axons by the combined signaling of nitric oxide and brain-derived neurotrophic factor. J. Neurosci. 20, 1458 – 1469. Finger, S., Almli, C.R., 1985. Brain damage and neuroplasticity: mechanisms of recovery or development? Brain Res. 357, 177 – 186. Fournier, A.E., Takizawa, B.T., Strittmatter, S.M., 2003. Rho kinase inhibition enhances axonal regeneration in the injured CNS. J. Neurosci. 23, 1416 – 1423. Gallo, G., Yee Jr., H.F., Letourneau, P.C., 2002. Actin turnover is required to prevent axon retraction driven by endogenous actomyosin contractility. J. Cell Biol. 158, 1219 – 1228. Gimenez y Ribotta, M., Gaviria, M., Menet, V., Privat, A., 2002. Strategies for regeneration and repair in spinal cord traumatic injury. Prog. Brain Res. 137, 191 – 212. Hill, C.E., Beattie, M.S., Bresnahan, J.C., 2001. Degeneration and sprouting of identified descending supraspinal axons after contusive spinal cord injury in the rat. Exp. Neurol. 171, 153 – 169. Houlem, J.D., Jin, Y., 2001. Chronically injured supraspinal neurons exhibit only modest axonal dieback in response to a cervical hemisection lesion. Exp. Neurol. 169, 208 – 217. Houle, J.D., Tessler, A., 2003. Repair of chronic spinal cord injury. Exp. Neurol. 182, 247 – 521. Jalink, K., van Corven, E.J., Hengeveld, T., Morii, N., Narumiya, S., Moolenaar, W.H., 1994. Inhibition of lysophosphatidate- and thrombininduced neurite retraction and neuronal cell rounding by ADP ribosylation of the small GTP-binding protein Rho. J. Cell Biol. 126, 801 – 810. Jian, X., Hidaka, H., Schmidt, J.T., 1994. Kinase requirement for retinal growth cone motility. J. Neurobiol. 25, 1310 – 1328. Jian, X., Szaro, B.G., Schmidt, J.T., 1996. Myosin light chain kinase: expression in neurons and upregulation during axon regeneration. J. Neurobiol. 31, 379 – 391. Kemp, B.E., Pearson, R.B., Guerriero Jr., V., Bagchi, I.C., Means, A.R., 1987. The calmodulin binding domain of chicken smooth muscle myosin light chain kinase contains a pseudosubstrate sequence. J. Biol. Chem. 262, 2542 – 2548. Lindgren, M., Hallbrink, M., Prochiantz, A., Langel, U., 2000. Cell-penetrating peptides. Trends Pharmacol. Sci. 21, 99 – 103. Luo, L., 2000. Rho GTPases in neuronal morphogenesis. Nat. Rev. Neurosci. 1, 173 – 180. McHale, M.K., Hall, G.F., Cohen, M.J., 1995. Early cytoskeletal changes following injury of giant spinal axons in the lamprey. J. Comp. Neurol. 353, 25 – 37. Oudega, M., Vargas, C.G., Weber, A.B., Kleitman, N., Bunge, M.B., 1999. Long-term effects of methylprednisolone following transection of adult rat spinal cord. Eur. J. Neurosci. 11, 2453 – 2464.
G. Gallo / Experimental Neurology 189 (2004) 112–121 Pallini, R., Fernandez, E., Sbriccoli, A., 1988. Retrograde degeneration of corticospinal axons following transection of the spinal cord in rats. A quantitative study with anterogradely transported horseradish peroxidase. J. Neurosurg. 68, 124 – 128. Pfitzer, G., 2001. Regulation of myosin phosphorylation in smooth muscle. J. Appl. Physiol. 91, 497 – 503. Pollack, E.D., Liebig, V., 1990. Stage-dependent spontaneous frog dorsal root ganglion neuritogenesis on polylysine in vitro. Anat. Rec. 228, 220 – 224. Raff, M.C., Whitmore, A.V., Finn, J.T., 2002. Axonal self-destruction and neurodegeneration. Science 296 (5569), 868 – 871 (May 3). Rochlin, M.W., Itoh, K., Adelstein, R.S., Bridgman, P.C., 1995. Localization of myosin II A and B isoforms in cultured neurons. J. Cell Sci. 108, 3661 – 3670. Ruchhoeft, M.L., Harris, W.A., 1997. Myosin functions in Xenopus retinal ganglion cell growth cone motility in vivo. J. Neurobiol. 32, 567 – 578. Schmidt, J.T., Morgan, P., Dowell, N., Leu, B., 2002. Myosin light chain phosphorylation and growth cone motility. J. Neurobiol. 52, 175 – 188. Spira, M.E., Oren, R., Dormann, A., Ilouz, N., Lev, S., 2001. Calcium, protease activation, and cytoskeleton remodeling underlie growth cone
121
formation and neuronal regeneration. Cell. Mol. Neurobiol. 21 (6), 591 – 604 (Dec.). Straight, A.F., Cheung, A., Limouze, J., Chen, I., Westwood, N.J., Sellers, J.R., Mitchison, T.J., 2003. Dissecting temporal and spatial control of cytokinesis with a myosin II Inhibitor. Science 299, 1743 – 1747. Wahl, S., Barth, H., Ciossek, T., Aktories, K., Mueller, B.K., 2000. EphrinA5 induces collapse of growth cones by activating Rho and Rho kinase. J. Cell Biol. 149, 263 – 270. Wylie, S.R., Chantler, P.D., 2003. Myosin IIA drives neurite retraction. Mol. Biol. Cell 14, 4654 – 4666. Wylie, S.R., Wu, P.J., Patel, H., Chantler, P.D., 1998. A conventional myosin motor drives neurite outgrowth. Proc. Natl. Acad. Sci. U. S. A. 95 (22), 12967 – 12972. Yarrow, J.C., Lechler, T., Li, R., Mitchison, T.J., 2003. Rapid de-localization of actin leading edge components with BDM treatment. BMC Cell Biol. 4, 5. Yoshizaki, H., Ohba, Y., Kurokawa, K., Itoh, R.E., Nakamura, T., Mochizuki, N., Nagashima, K., Matsuda, M., 2003. Activity of Rho-family GTPases during cell division as visualized with FRET-based probes. J. Cell Biol. 162 (2), 223 – 232 (Jul. 21).