Axotomy inhibits the slow axonal transport of tubulin in the squid giant axon.
To test this possibility, the lateral giant axon was transected, and tubulin transport was then assayed. For each experiment a squid (Loliguncula brevis, obtained most months of the year from the National Resource Center for Cephalopods, Galveston TX, or Loligo pealeii, obtained during May or October from the Marine Resource Center. Marine Biological Laboratory, Woods Hole, MA) was first anesthetized with 1% ethanol in seawater for about 5 rain. The stellate ganglia were then located by eye through the mantle opening. A pair of Vannas micro-dissection scissors angled on the flat was inserted through the mantle opening, and one of the first lateral giant axons was transected (axotomized). The squid was then returned to a fresh seawater tank for 2 days. Squid were usually fed once a day for 2 days after return to the seawater tanks. After 2 days the squid was sacrificed, and the distal end of the transected lateral axon as well as the uncut control axon from the other side were removed, cleaned, and prepared for the slow transport assays. The air pressure system described previously (2) was used to inject both fluorescently labeled tubulin (Cytoskeleton, Inc.) and a marker oil drop into the control and transected axons. Axons were injected in the middle of the 2-4 cm isolated axons. Transport was visualized by recording the fluorescent image over time with a Zeiss microscope equipped with a Photometrics Cool Snap HQ camera driven by Openlab software (Improvision).
In the control axons (n = 6), tubulin diffused while simultaneously being transported anterogradely (to the right in Fig. 1A). The injection site is marked by an arrowhead. As illustrated in Figure 1B, this anterograde tubulin transport was arrested 2 days after axotomy. The tubulin diffused in both directions away from the injection site (arrowhead) but was not transported anterogradely (n = 6).
[FIGURE 1 OMITTED]
To determine whether the inhibition of tubulin transport after axotomy might be due to some nonspecific damage or degeneration of the isolated axon. a number of structural and biochemical tests of axonal health were performed. We have previously demonstrated that when a squid axon is damaged or becomes leaky, its light scattering increases, mitochondria become swollen, and its cytoplasm becomes disorganized (5). Though most of these signs of damage could be seen at distances closer than 1 mm from the transaction site (not shown), the rest of the 1-4 cm axons showed no signs of axonal damage. As illustrated in Figure 1 (compare C with D), no change in darkfield light scattering was detected in these or other transected axons (n = 12); and as illustrated in Figure 1 (compare E with F), the cytoskeletal elements appeared normal in the transected axon, and the mitochondria were not swollen (n = 10).
Since protein loss in general, and neurofilament proteolysis in particular (6), are sensitive indicators of axonal damage in squid as well as in other axons, we compared the levels of neurofilament and other axonal proteins in transected and control axons. Total protein was measured as follows: Equal lengths (from 1-4 cm in different squids) of an axotomized axon and its paired control were excised in calcium-free seawater. The isolated axon was then dissolved in solubilization buffer (2% SDS, 2% BME, 8 M urea, Tris pH 6.8), and the solubilized proteins were separated by 7.5% PAGE. To measure changes in total axonal proteins, the separated proteins were stained with Ruby protein stain (Molecular Probes). A dilution series of control axons was run with each experiment to verify that optical density was linearly related to protein concentration in each run (not shown). As illustrated in Figure 1G, axotomy produced no changes in neurofilament (NF200 and NF60), tubulin, or any other major axonal protein. Equal lengths of control (1c) and transected (1t) axons from the same squid (squid 1) had virtually identical protein-staining profiles. Smaller paired axons from a smaller squid (squid 2) had proportionately less protein (2c and 20, but there was no visible difference in protein levels between the control axon (2c) and the previously transected axon (2t) in this or other squids tested (n = 6).
To determine if the loss of conventional kinesin might be responsible for the loss of slow tubulin transport (3), we measured kinesin levels in the control and transected axons. Since conventional kinesin and other potential motors are minor constituents of axoplasm, Ruby protein stain would not detect these protein. To measure the kinesin levels, axons were cut, cleaned, dissolved, and run on PAGE as above. The PAGE-separated proteins were then transferred onto PVDF paper by western blotting in Tris-glycine buffer, and exposed to an anti-kinesin antibody (3) (Chemicon MAB 1614). The amount of antibody was detected with chemiluminescence. As with the protein measurements, axoplasm standards were run to insure that the recorded optical densities varied linearly with kinesin concentration. As illustrated in Figure 1H (Kinesin), there was no significant difference in kinesin levels between the control and the transected axon in either squid 1 (compare 1c with 1t) or squid 2 (compare 2e with 2t) or with any other animals tested (n = 9).
The present experiment suggests that some unidentified factor essential to axonal tubulin transport is lost or inhibited 2 days after axotomy. This inhibition is not due to the depletion of conventional kinesin. It could be due to the inactivation of conventional kinesin or to the inactivation or depletion of some other motor or factor essential to axonal tubulin transport. This model of protein transport in the squid giant axon may facilitate the identification of some of the factors essential to the control and maintenance of slow axonal transport.
The author thanks James Galbraith and Thomas Reese for helpful discussions and comments.
(1.) Terasaki, M., A. Schmidek, J. A. Galbraith, P. E. Gallant, and T. S. Reese. 1995. Proc. Natl. Acad. Sci. USA 92: 11,500-11.503.
(2.) Galbraith, J. A., T. S. Reese, M. L. Schlief, and P. E. Gallant. 1999. Proc. Natl. Acad. Sci. USA 96:11,589-11,594.
(3.) Terada, S., M. Kinjo, and N. Hirokawa. 2000. Cell 103:141 155.
(4.) Gallant, P. E. 2000. J. Neurocytol. 29: 779-782.
(5.) Gallant, P. E., and J. A. Galbraith. 1997. J. Neurotrauma 14: 811-822.
(6.) Gallant, P. E., H. C. Pant, R. M. Pruss, and H. Gainer. 1986. J. Neurochem. 46: 1573-1581.
P. E. Gallant
National Institutes of Health, Bethesda, MD
Marine Biological Laboratory, Woods Hole, MA
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|Publication:||The Biological Bulletin|
|Date:||Oct 1, 2003|
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