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15.9: Video- The Unfixed Spinal Cord - Biology

15.9: Video- The Unfixed Spinal Cord - Biology


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15.9: Video- The Unfixed Spinal Cord

Neuroanatomy Video Lab: Brain Dissections

This series of Neuroanatomy “lessons” has two principal objectives. The first is to provide viewers access to human brain specimens, something lacking in many places. The second is to simplify the anatomy, omitting many details, and making many generalizations. The reason for this is to keep the focus on localization of the patient’s disease within the nervous system. Students are often overwhelmed with excessive detail making correlation of structure and function more difficult. The presentation sequence is in a logical order for a class, but each video is designed to standalone and be used just-in-time to make a clinical point.

The series originated when teaching with a neurologist in Ghana and Kenya where it was clear that anatomy taught separated from the clinic by many years was not retained. Here I present just enough anatomy to localize the patient’s problem. These videos can be used before or after a clinical case presentation. Apologies are offered for a few minor errors that could not be corrected due to the one time recording opportunity as well as an unscripted personal style. The audience for these videos can be nurses, physician assistants, medical students, residents, registrars, house officers or anyone interested in the brain.

Videos can also be used in conjunction with the NeurologicExam Website → All of the videos have English captions to assist those for whom English is not their native language or to visualize the many new terms.


Editorial Reviews

Review

"This atlas provides an excellent, detailed map of the entire spinal cord of both rat and mouse. The photomicrographs are outstanding, the labelling is clear and the illustrations should serve as outstanding examples of what high quality staining and immunocytochemistry should look like. This information has not been available in any atlas of the CNS before, and will be an extremely useful resource for all neuroscientist interested in this part of the nervous system and a 'must-have' for spinal cord labs."

Jacqueline C. Bresnahan, Professor, Department of Neurological Surgery, Brain and Spinal Injury Center, University of California at San Francisco, USA

"The Spinal Cord is an authoritative and detailed account of the development, organization and function of the spinal cord. Written by a series of experts, the book contains enlightening chapters that cover the anatomy and the architecture of the spinal cord in a clear and logical fashion. Attention to special topics, such as spinal cord injury and micturition, is unprecedented and unusually informative. The comprehensive atlas, along with the diagrams and list of references, will be of considerable use to the students of the nervous system, as well as the most senior of investigators. It is an excellent volume!"

Moses V. Chao, Professor of Cell Biology, Physiology and Neuroscience and Psychiatry, Molecular Neurobiology Program, Skirball Institute of Biomolecular Medicine, NYU School of Medicine, New York, USA

Review

About the Author

Charles Watson is a neuroscientist and public health physician. His qualifications included a medical degree (MBBS) and two research doctorates (MD and DSc). He is Professor Emeritus at Curtin University, and holds adjunct professorial research positions at the University of New South Wales, the University of Queensland, and the University of Western Australia.
He has published over 100 refereed journal articles and 40 book chapters, and has co-authored over 25 books on brain and spinal cord anatomy. The Paxinos Watson rat brain atlas has been cited over 80,000 times. His current research is focused on the comparative anatomy of the hippocampus and the claustrum.
He was awarded the degree of Doctor of Science by the University of Sydney in 2012 and received the Distinguished Achievement Award of the Australasian Society for Neuroscience in 2018.

Professor George Paxinos, AO (BA, MA, PhD, DSc) completed his BA at The University of California at Berkeley, his PhD at McGill University, and spent a postdoctoral year at Yale University. He is the author of almost 50 books on the structure of the brain of humans and experimental animals, including The Rat Brain in Stereotaxic Coordinates, now in its 7th Edition, which is ranked by Thomson ISI as one of the 50 most cited items in the Web of Science. Dr. Paxinos paved the way for future neuroscience research by being the first to produce a three-dimensional (stereotaxic) framework for placement of electrodes and injections in the brain of experimental animals, which is now used as an international standard. He was a member of the first International Consortium for Brain Mapping, a UCLA based consortium that received the top ranking and was funded by the NIMH led Human Brain Project. Dr. Paxinos has been honored with more than nine distinguished awards throughout his years of research, including: The Warner Brown Memorial Prize (University of California at Berkeley, 1968), The Walter Burfitt Prize (1992), The Award for Excellence in Publishing in Medical Science (Assoc Amer Publishers, 1999), The Ramaciotti Medal for Excellence in Biomedical Research (2001), The Alexander von Humbolt Foundation Prize (Germany 2004), and more. --This text refers to the hardcover edition.


  • Editorial &rlm : &lrm Academic Press Edición Illustrated (12 noviembre 2008)
  • Idioma &rlm : &lrm Inglés
  • Pasta dura &rlm : &lrm 387 páginas
  • ISBN-10 &rlm : &lrm 0123742471
  • ISBN-13 &rlm : &lrm 978-0123742476
  • Dimensiones &rlm : &lrm 21.84 x 2.79 x 27.94 cm

Críticas

This atlas provides an excellent, detailed map of the entire spinal cord of both rat and mouse. The photomicrographs are outstanding, the labelling is clear and the illustrations should serve as outstanding examples of what high quality staining and immunocytochemistry should look like. This information has not been available in any atlas of the CNS before, and will be an extremely useful resource for all neuroscientist interested in this part of the nervous system and a 'must-have' for spinal cord labs. --Jacqueline C. Bresnahan, Professor, Department of Neurological Surgery, Brain and Spinal Injury Center, University of California at San Francisco, USA

The Spinal Cord is an authoritative and detailed account of the development, organization and function of the spinal cord. Written by a series of experts, the book contains enlightening chapters that cover the anatomy and the architecture of the spinal cord in a clear and logical fashion. Attention to special topics, such as spinal cord injury and micturition, is unprecedented and unusually informative. The comprehensive atlas, along with the diagrams and list of references, will be of considerable use to the students of the nervous system, as well as the most senior of investigators. It is an excellent volume! --Moses V. Chao, Professor of Cell Biology, Physiology and Neuroscience and Psychiatry, Molecular Neurobiology Program, Skirball Institute of Biomolecular Medicine, NYU School of Medicine, New York, USA

Biograf໚ del autor

Charles Watson is a neuroscientist and public health physician. His qualifications included a medical degree (MBBS) and two research doctorates (MD and DSc). He is Professor Emeritus at Curtin University, and holds adjunct professorial research positions at the University of New South Wales, the University of Queensland, and the University of Western Australia.
He has published over 100 refereed journal articles and 40 book chapters, and has co-authored over 25 books on brain and spinal cord anatomy. The Paxinos Watson rat brain atlas has been cited over 80,000 times. His current research is focused on the comparative anatomy of the hippocampus and the claustrum.
He was awarded the degree of Doctor of Science by the University of Sydney in 2012 and received the Distinguished Achievement Award of the Australasian Society for Neuroscience in 2018.

Professor George Paxinos, AO (BA, MA, PhD, DSc) completed his BA at The University of California at Berkeley, his PhD at McGill University, and spent a postdoctoral year at Yale University. He is the author of almost 50 books on the structure of the brain of humans and experimental animals, including The Rat Brain in Stereotaxic Coordinates, now in its 7th Edition, which is ranked by Thomson ISI as one of the 50 most cited items in the Web of Science. Dr. Paxinos paved the way for future neuroscience research by being the first to produce a three-dimensional (stereotaxic) framework for placement of electrodes and injections in the brain of experimental animals, which is now used as an international standard. He was a member of the first International Consortium for Brain Mapping, a UCLA based consortium that received the top ranking and was funded by the NIMH led Human Brain Project. Dr. Paxinos has been honored with more than nine distinguished awards throughout his years of research, including: The Warner Brown Memorial Prize (University of California at Berkeley, 1968), The Walter Burfitt Prize (1992), The Award for Excellence in Publishing in Medical Science (Assoc Amer Publishers, 1999), The Ramaciotti Medal for Excellence in Biomedical Research (2001), The Alexander von Humbolt Foundation Prize (Germany 2004), and more.


Postnatal murine spinal cord represents a good model system to study mammalian central nervous system myelination in vivo as a basis for further studies in demyelinating diseases.

Transcriptional changes were analyzed in SJL/J mice on postnatal day 0, 14, 49 and 231 (P0, P14, P49, P231) employing Affymetrix GeneChip Mouse Genome 430 2.0 Arrays. Additionally, marker gene signatures for astrocyte and oligodendrocyte lineage-stages were defined to study their gene expression in more detail. In addition, immunohistochemistry was used to quantify the abundance of commonly used glial cell markers.

6092 differentially regulated genes (DEGs) were identified. The up-regulated DEGs at P14, P49 and P231 compared to P0 exhibited significantly enriched associations to gene ontology terms such as myelination and lipid metabolic transport and down-regulated DEGs to neurogenesis and axonogenesis. Expression values of marker gene signatures for neural stem cells, oligodendrocyte precursor cells, and developing astrocytes were constantly decreasing, whereas myelinating oligodendrocyte and mature astrocyte markers showed a steady increase. Molecular findings were substantiated by immunohistochemical observations.

The transcriptional changes observed are an important reference for future analysis of degenerative and inflammatory conditions in the spinal cord.


Results

Kv3.1b is localized to the nodes of Ranvier in the CNS

The Kv3.1 gene gives rise to two distinct channel subunits, Kv3.1a and Kv3.1b, by alternative splicing (Luneau et al., 1991). Their predicted protein sequences diverge only at their C terminals. We focused on Kv3.1b because it is far more abundant than Kv3.1a in the adult rat brain (Perney et al., 1992 Gan and Kaczmarek, 1998). Our preliminary investigations revealed that cryosections of unfixed spinal cord have stronger Kv3.1b staining than do tissues fixed with either 4% paraformaldehyde or Zamboni's fixative, and that the gray matter was strongly stained. To localize the nodes of Ranvier, we stained transverse sections of unfixed adult mouse spinal cord with a rabbit antiserum against Kv3.1b or Kv3.2 and a mouse monoclonal antibody that recognizes a sequence common to all Nav channel α subunits. Weak Kv3.2 staining was seen in the spinal cord gray matter, but not in the white matter (data not shown). As shown in Figure 1A𠄿 , numerous nodes were labeled for Kv3.1b in all white matter tracts, but the proportion of Kv3.1b-positive nodes differed ( Fig. 2A ). In the lateral column ( Fig. 1A,B ) and the ventral funiculus ( Fig. 1C,D ), nearly all nodes were colabeled for Nav and Kv3.1b channels. In contrast, in the corticospinal tract ( Fig. 1E,F ), relatively fewer nodes were labeled for Kv3.1b. When observed at high magnification, Kv3.1b and Nav channels staining appeared circular, confirming that they were expressed at nodes ( Fig. 3A𠄼 , insets). Moreover, in longitudinal sections, Kv3.1b staining precisely matched that of Nav channels ( Fig. 3A𠄼 , double arrowheads). The staining for Kv3.1b was specific because it was abolished by preincubating the antiserum with the peptide (data not shown). Furthermore, no Kv3.1b staining was found in Kv3.1-deficient mice ( Fig. 1G,H ), which lack both Kv3.1a and Kv3.1b isoforms (Ozaita et al., 2002).

Localization of Kv3.1b in the adult mouse spinal cord. Transverse sections of unfixed thoracic spinal cord were double labeled with a rabbit Kv3.1b antiserum [tetramethylrhodamine isothiocyanate (TRITC)] and a “pan” monoclonal antibody (FITC) against Nav channels (panNav). In the lateral (A, B) and ventral (C, D) columns of the spinal cord, the majority of the nodes were double labeled, but in the dorsal column (dc) and especially in the corticospinal tract (cst) (E, F), only a subset of nodes were Kv3.1b-positive. Note the intense staining for Kv3.1b in the dorsal horn (dh) and ventral horn (vh) of the spinal cord. In Kv3.1-deficient (-/-) mouse, no labeling was observed for Kv3.1b (the ventral column is shown) (G, H). Scale bar, 20 μm.

Kv3.1b labeling in adult mouse spinal cord, optic nerve, and DRG. Longitudinal sections of the ventral column (A𠄼) and dorsal horn (G–I) of the spinal cord, as well as optic nerve (D𠄿) and DRG (J–L), double-labeled with a rabbit Kv3.1b antiserum (TRITC) and a monoclonal antibody (FITC) against Nav channels (panNav) are shown. In A𠄼, Kv3.1b is colocalized exclusively with Nav channels in the nodes (double arrowheads). The inset shows a node in a transverse section. In D𠄿, few nodes were Kv3.1b-positive. In G–I, Kv3.1b appears to be expressed, at least in part, in the somatic membrane of neurons, but was not colocalized with Nav channels in the initial segments (arrows). In J–L, many somata expressed Nav channels, but few expressed Kv3.1b. Arrows indicate initial segments. Scale bars: (in C) A𠄼, 20 μm (in F) D𠄿, 20 μm (in I) G–I, 20 μm (in L) J–L, 20 μm (in C, inset) A𠄼, insets, 5 μm.

Relationship between Kv3.1b expression and nodal diameter in the dorsal and ventral funiculus of adult mouse spinal cord. The number of nodes and Kv3.1b-positive nodes are represented in function of the nodal diameter range for the dorsal (C) and ventral (D) funiculus. In B, the percentage of Kv3.1b-positive nodes in the whole spinal cord are represented in the function of the nodal diameter range. Note that almost all large nodes (ϡ.8 μm) are Kv3.1b-positive, whereas 7�% of the small nodes (ρ.8 μm) are Kv3.1b-negative, and that Kv3.1b is more represented in small nodes of the ventral funiculus than in the dorsal funiculus. Data were obtained from two mice and are represented as means ± SD n = 1394.

We suspected that the reason for the variation in nodal staining was the variation in the composition of axons in different tracts. In particular, the size of myelinated axons appeared to correlate with Kv3.1b nodal staining. The CST contains many small myelinated axons (Hildebrand, 1972 Murray and Blakemore, 1980 Chung and Coggeshall, 1987), whereas the ventral and lateral funiculi contain many large myelinated axons. To address this issue, we immunostained sections of mouse and rat optic nerves, which mainly contain small myelinated fibers (Matheson, 1970), and found that 14% of the nodes were labeled for Kv3.1b. The nodes that were stained appeared to have a larger diameter than the unlabeled nodes ( Fig. 3D𠄿 ). To confirm that Kv3.1b expression is related to the diameter of the fibers, a quantitative study was performed, and the number of nodes and Kv3.1b-positive nodes relative to the diameter range of the nodes was measured. We found that almost every node with a diameter ϡ.8 mm was Kv3.1b-positive, whereas 54 to 93% of the nodes having a diameter ρ.8 mm were Kv3.1b-positive ( Fig. 2B ). These data support a relationship between the axonal diameter and the nodal localization of Kv3.1b. However, because small nodes in the ventral funiculus are more often Kv3.1b-positive than those in the dorsal funiculus ( Fig. 3C,D ), it appears that other factors also influence the expression of Kv3.1b.

Because nodes and initial segments both have ankyrin G and Nav channels, we determined whether Kv3.1b was expressed in initial axonal segments by labeling longitudinal sections of the spinal cord as well as sections of dorsal root ganglia. Although the gray matter was intensely labeled (likely corresponding to surface staining of somata and presynaptic terminals) (Sekirnjak et al., 1997), initial segments were not stained ( Fig. 3G–I ). The sensory neurons in the dorsal ganglia, in contrast, had little Kv3.1b staining this may be related to the absence of synapses on their cell bodies ( Fig. 3J–L ). As in the spinal cord, the initial segments of sensory neurons were Kv3.1b-negative.

Localization of Kv3.1b and Kv3.2 in the PNS

We examined the localization of Kv3.1b and Kv3.2 in teased fibers from adult mouse sciatic nerves. PNS nodes were Kv3.2-negative (data not shown), and only 23% of the nodes expressed Kv3.1b ( Fig. 4A,B ) the labeling even in these nodes was less intense than that in the CNS. When it was possible to follow single fibers, we observed that all of the nodes were either stained or unstained, suggesting that Kv3.1b was expressed in a specific subset of fibers, perhaps related to their function. To elucidate this point, we stained teased fibers from both the dorsal and the ventral roots, but found Kv3.1-positive nodes in both small- and large-diameter fibers of the dorsal and ventral roots (data not shown). We also examined the development of Kv3.1b expression in the dorsal and ventral roots. At P4 and P8, no nodal staining was observed some clusters were observed at P12, and many more at P15 and P21 (data not shown). These observations exclude the possibility that PNS nodes lose expression of Kv3.1 during development. Thus, the functional significance of Kv3.1b-positive PNS nodes remains to be established.

Few PNS nodes express Kv3.1b. Unfixed teased fibers from adult mouse sciatic nerve were double labeled with a rabbit Kv3.1b antiserum (TRITC,A) and a monoclonal antibody against Nav channels (FITC, B). One Kv3.1b-positive node is indicated (double arrowheads) two other nodes are Kv3.1b-negative. Scale bar, 10 μm.

Because motor and sensory neurons have myelinated fibers with both CNS and PNS myelin, we examined sections of the ventral root entry zone, the transition zone between CNS and PNS myelin for motor fibers. Sections colabeled for Kv3.1b and Kv1.2 show that Kv3.1b-positive nodes are found in the spinal cord but not in the roots, whereas Kv1.2-positive juxtaparanodes were found in both places ( Fig. 5A𠄼 ). Thus, the strong nodal labeling of Kv3.1b in motor axons appears to depend on some aspect of the CNS environment, presumably related to either oligodendrocytes or astrocytes.

Differential localization of Kv3.1b and Kv1.2 in CNS axons. These are images of longitudinal sections of the ventral column (vc) of unfixed lumbar spinal cord from adult mice. In the spinal cord, the narrow band of nodal Kv3.1b staining (TRITC A) is flanked by two broader regions of juxtaparanodal Kv1.2 staining (FITC B). C, Merge. Note the relative lack of labeling for Kv3.1b in the PNS nodes (double arrowheads) of the ventral rootlet (vr). Scale bar, 10 μm.

Developmental expression of Kv3.1b

Ankyrin G is the first molecular component that becomes localized in developing CNS nodes Nav channels, L1, neurofascin and βIV-spectrin are recruited later (Jenkins and Bennett, 2001). To determine when Kv3.1b becomes localized to nodes, we immunostained transverse sections of the thoracic spinal cord from rats at P4, P8, P12, and P21. At P4, there were clusters of Nav channels in the ventral and lateral columns ( Fig. 6B ), but no Kv3.1b staining in the white matter or even the gray matter ( Fig. 6A ). At P8, there was weak Kv3.1b staining in the gray matter, and clusters of Kv3.1b in a minority of nodes in the ventral column ( Fig. 6C,D ). In longitudinal sections of the ventral column, Kv3.1b seemed to be selectively localized to “mature” nodes (short nodes rather than in elongated ones Rasband et al., 1999a). At P12, the number, as well as the staining intensity of Kv3.1b-positive nodes increased markedly ( Fig. 6E,F ), and in longitudinal sections, Kv3.1b clusters appeared mostly in short, mature-appearing nodes ( Fig. 7A ). At P21, the distribution of Kv3.1b-positive nodes ( Fig. 6G,H ) appeared similar to that in adult rats.

Delayed appearance of Kv3.1b at CNS nodes. These are images shown as negatives (the nodes are dark) of transverse sections of unfixed thoracic spinal cord from P4 (A, B) P8 (C, D), P12 (E, F), and P21 (G, H) rats. The ventral column (vc) is shown, as well as the ventral horn (vh) in A𠄿. At P4, Nav channels but not Kv3.1b channels were clustered. Kv3.1b clusters were present by P8, and their number, as well as that of Nav channels, increased from P4 to P21. At P21, nodes were labeled for both Nav channels and Kv3.1b. Scale bars: (in B) A, B, 20 μm (in H) B–H, 20 μm.

Kv3.1b appears before Kv1.2 in CNS myelinated fibers. These are longitudinal sections of the ventral column of unfixed rat ventral lumbar spinal cords at P12 (A), P15 (B), and P21 (C). At P12 and P15, nodal Kv3.1b staining was often not associated with juxtaparanodal Kv1.2 staining, whereas at P21 most nodes were flanked by juxtaparanodal Kv1.2 labeling. Scale bars, 10 μm.

We considered the possibility that Kv3.1b might cluster concomitantly with Kv1.1/Kv1.2 subunits. Kv1.1 and Kv1.2 are the last channels to be segregated in developing CNS (Rasband et al., 1999b), at approximately P8 in the spinal cord (Wang et al., 1995), and their clustering in the juxtaparanodal region depends on the formation of normal myelin sheaths, especially intact septate-like junctions (Baba et al., 1999 Mathis et al., 2001 Arroyo et al., 2002). Longitudinal sections through the ventral columns of lumbar spinal cords of P4, P8, P12, P15, and P21 rats were double labeled for Kv3.1b and Kv1.2. No staining for Kv1.2 was observed at P4 in the spinal cord, and few clusters of Kv1.2 were observed at P8 (data not shown). At P12, most Kv3.1b-positive nodes had no associated Kv1.2 staining ( Fig. 7A ). At P15, the proportion of Kv3.1b-positive nodes associated with Kv1.2 staining increased, and by P21 all Kv3.1b-positive nodes were flanked by juxtaparanodal Kv1.2 staining ( Fig. 7B,C ). Together, these results show that ion channel clustering in myelinated fibers follow the order: Nav channels, then Kv3.1b subunits, then Kv1.2 subunits.

Does nodal clustering of Kv3.1b depend on myelination?

To answer this question, we studied the localization of Kv3.1b in md rats, which typically die at approximately P21 from severe dysmyelination associated with oligodendrocyte cell death (Grinspan et al., 1998). Even at this age, few axons are myelinated in the spinal cord axonal segments are ensheathed by few wraps of oligodendrocyte processes, which can be detected by their expression of MAG, or are totally devoid of ensheathment (Dentinger et al., 1982 Rosenbluth, 1987 Arroyo et al., 2002). Nevertheless, their spinal cords contain normal numbers of node-like clusters of Nav channels and ankyrin G, even in regions devoid of oligodendrocytes (Arroyo et al., 2002). As shown in Figure 8 , Kv3.1b and Nav channels were colocalized in the spinal cord of P21 md rats (A𠄼) and their age-matched WT littermates (D𠄿). Double labeling for MAG demonstrated that some node-like clusters of Kv3.1b were flanked by MAG-ensheathed segments, but many were found in unensheathed axonal segments ( Fig. 8G ). In contrast, the localization of the Kv1.2 subunits was highly altered in md rats: Kv1.2 subunits abutted the node-like clusters of Kv3.1b ( Fig. 8H ) instead of being concentrated in the juxtaparanodal regions as in WT rats ( Fig. 8I ). These results show that node-like clusters of Kv3.1b behave similarly to clusters of Nav channels in md rats (Arroyo et al., 2002): both are formed during development and maintained at least temporarily even in the absence of myelin sheaths.

Node-like clusters of Kv3.1b in md rats. Longitudinal sections through the ventral funiculus of unfixed lumbar spinal cord from P21 md rats and their WT male littermates were double labeled with a rabbit Kv3.1b antiserum (TRITC) and a monoclonal antibody (FITC) against Nav channels (pan Nav A𠄿), MAG (G), or Kv1.2 (H, I). In A𠄿, note that in both md and WT rats, node-like clusters Kv3.1b colocalized with Nav channels (double arrows). In G, note that many node-like clusters of Kv3.1b (arrowheads) were found in regions devoid of MAG-positive oligodendrocyte processes. The asterisks mark oligodendrocyte nuclei. In H, I, note that the localization of Kv1.2 (arrows) was altered in md rats, abutting the node-like clusters of Kv3.1b (double arrows). Scale bars: (in C) A𠄼, 10 μm (in F) D𠄿, 10 μm G, 10 μm H, 10 μm I, 10 μm.

Kv3.1b is associated with ankyrin G in brain membranes

The colocalization of Kv3.1b, Nav channels, and ankyrin G in CNS nodes suggested that Kv3.1b may interact directly or indirectly with ankyrin G. The nodal isoforms of ankyrin G interact with several nodal proteins such as Nav channel α and β subunits, neurofascin, and neural-glial-related CAM (NrCAM) (Malhotra et al., 2000 Bennett and Chen, 2001). To determine the level of expression of Kv3.1b and Kv3.2 in the brain, spinal cord, optic nerve, and sciatic nerve, we performed a Western blot analysis of these tissues ( Fig. 9 ). Rat brain and spinal cord expressed more Kv3.1b than did optic nerve Kv3.1b could not be detected in rat sciatic nerve the above data are in keeping with the finding that CNS nodes have more robust Kv3.1b expression than do PNS nodes. The anti-Kv3.1b antiserum recognized a diffuse band of �� kDa, significantly greater than the 66 kDa expected for the Kv3.1b polypeptide obtained by translation of the Kv3.1b cDNA (Luneau et al., 1991), but in good agreement with the results of Weiser et al. (1995) on rat brain membranes. This band is almost certainly Kv3.1b, because it is absent in brain membranes from Kv3.1b-deficient mice ( Fig. 9A ). Kv3.2 was also detected in brain membranes, and to a lesser extent in the spinal cord, with a molecular weight range of 75� kDa ( Fig. 9B ). However, Kv3.2 was not detected in optic and sciatic nerves, in agreement with the immunostaining. We did not detect alterations in Kv3.2 staining in Kv3.1-deficient mouse spinal cord or brain in particular, nodes were not Kv3.2-positive (data not shown). In addition, we did not detect any alterations in the labeling for contactin, Caspr, Kv1.1, ankyrin G or Nav channels (data not shown), suggesting that the absence of Kv3.1 does not alter the organization of these axonal proteins in myelinated fibers.

Distribution and interaction with ankyrin G of Kv3.1b and Kv3.2 subunits. A, B, Membrane proteins (100 μg) from Kv3.1-deficient mouse brain and rat brain, spinal cord, optic nerve, and sciatic nerve were fractioned in SDS gel and immunoblotted for Kv3.1b (A) and Kv3.2 (B). Kv3.1b was expressed in rat brain and spinal cord, and to a lesser extents in the optic nerve. No band was observed in Kv3.1-deficient mouse. Kv3.2 was detected in the mouse and rat brain and to a lesser extent in the rat spinal cord. C, Brain and spinal cord membranes (200 μg) were immunoprecipitated separately for Kv3.1b or Kv3.2 and then immunoblotted for Kv3.1b. Note that Kv3.1b coimmunoprecipitates extensively with Kv3.2 in the brain but not in the spinal cord. D, E, Brain and spinal cord membranes (200 μg) were immunoprecipitated with rabbit anti-ankyrin G (Ank-G), rabbit anti-Kv3.1b, rabbit anti-Kv3.2, or mouse anti-Kv1.2 antibodies, then immunoblotted for Kv3.1b (D) or ankyrin G (E). Brain membrane (BM) was used as a positive control. Kv3.1b coimmunoprecipitates with ankyrin G and Kv3.2, but not Kv1.2 (D). A low-molecular-weight isoform of ankyrin G was pulled down with Kv3.1b, but not with either Kv3.2 or Kv1.2 (E). Molecular weight markers are shown on the right in kilodaltons.

Because subunits of the voltage-dependent K + channels can form heteromeric channels with members of their own family (Kv1, Kv2, Kv3, and Kv4), we immunoprecipitated Kv3.2 to determine whether Kv3.1b and Kv3.2 subunits could interact. When Kv3.2 was precipitated, it pulled down more Kv3.1b in brain than in the spinal cord ( Fig. 9C ), in keeping with the relative lack of Kv3.2 in the spinal cord. These results indicate that Kv3.1b could specifically form heteromeric channels with Kv3.2 in the CNS, but if they do, it is not at nodes.

To determine whether ankyrin G interacts with Kv3.1b, CNS membranes were incubated with antisera against ankyrin G or Kv3.1b, and the immunoprecipitates were probed for the same proteins. We used immunoprecipitates for Kv3.2 as a positive control and immunoprecipitates for Kv1.2 as a negative control. When ankyrin G was precipitated from brain membranes, it pulled down Kv3.1b ( Fig. 9D ). However, Kv3.1b did not pull down high-molecular-weight isoforms of ankyrin G, but pulled down an isoform of ankyrin G of � kDa ( Fig. 9E ). Neither Kv3.2 nor Kv1.2 coimmunoprecipitated this isoform of ankyrin G ( Fig. 9E ). Because ankyrin G is sensitive to proteolysis (Davis and Bennett, 1984), this isoform could correspond to a degradation product of the high-molecular-weight nodal isoforms of ankyrin G. Alternatively, this result could indicate that an isoform of ankyrin G that is not localized to nodes interacts with Kv3.1b, because both ankyrin G and Kv3.1b are present in gray matter.

Does Kv3.1b participate with AP repolarization in CNS nodes?

To answer this question, we tested the effects of 4-AP, a potent blocker of Kv3.1b (Grissmer et al., 1994), on the CAPs recorded from sciatic nerve, spinal cord ventral funiculus, and optic nerve of adult rats. 4-AP slightly increased the duration of the CAPs of sciatic nerves ( Fig. 10A ), but broadened the CAPs recorded from the spinal cord ventral funiculus and optic nerve ( Fig. 10A ). The lack of effect of 4-AP in the PNS could be attributable to the weak expression of Kv3.1b in sciatic nerve, but 4-AP affected the optic nerve, in which few nodes were Kv3.1b-positive. To resolve this discrepancy, we recorded CAPs from the optic nerves of Kv3.1-deficient and age-matched WT mice. CAPs recorded from WT and Kv3.1-deficient mice had similar onsets, peaks, and duration ( Fig. 10B ). The recruitment and refractory period of the fibers were also similar (data not shown). Moreover, 4-AP broadened the CAPs from both the WT and the mutant mice ( Fig. 10B ). These results suggest that the effects of 4-AP in the optic nerve axons are not mediated by Kv3.1 another nodal K + channel subunit could mediate these effects.

Effects of 4-AP on the CAPs of rat sciatic nerves, spinal cord ventral funiculi and optic nerves, as well as WT and Kv3.1-deficient mouse optic nerves. A, 4-AP (500 μ m ) increased slightly the duration of the CAPs from rat sciatic nerves (top traces, n = 2), but prolonged greatly the CAPs from the ventral funiculus (middle traces, n = 5) and optic nerve (bottom traces, n = 6).B, In WT (n = 2) and Kv3.1-deficient mice (n = 2), the characteristics of the optic nerve CAPs were identical, and 4-AP had similar effects.


Review

"This atlas provides an excellent, detailed map of the entire spinal cord of both rat and mouse. The photomicrographs are outstanding, the labelling is clear and the illustrations should serve as outstanding examples of what high quality staining and immunocytochemistry should look like. This information has not been available in any atlas of the CNS before, and will be an extremely useful resource for all neuroscientist interested in this part of the nervous system and a 'must-have' for spinal cord labs." --Jacqueline C. Bresnahan, Professor, Department of Neurological Surgery, Brain and Spinal Injury Center, University of California at San Francisco, USA

"The Spinal Cord is an authoritative and detailed account of the development, organization and function of the spinal cord. Written by a series of experts, the book contains enlightening chapters that cover the anatomy and the architecture of the spinal cord in a clear and logical fashion. Attention to special topics, such as spinal cord injury and micturition, is unprecedented and unusually informative. The comprehensive atlas, along with the diagrams and list of references, will be of considerable use to the students of the nervous system, as well as the most senior of investigators. It is an excellent volume!" --Moses V. Chao, Professor of Cell Biology, Physiology and Neuroscience and Psychiatry, Molecular Neurobiology Program, Skirball Institute of Biomolecular Medicine, NYU School of Medicine, New York, USA

About the Author

Charles Watson is a neuroscientist and public health physician. His qualifications included a medical degree (MBBS) and two research doctorates (MD and DSc). He is Professor Emeritus at Curtin University, and holds adjunct professorial research positions at the University of New South Wales, the University of Queensland, and the University of Western Australia.
He has published over 100 refereed journal articles and 40 book chapters, and has co-authored over 25 books on brain and spinal cord anatomy. The Paxinos Watson rat brain atlas has been cited over 80,000 times. His current research is focused on the comparative anatomy of the hippocampus and the claustrum.
He was awarded the degree of Doctor of Science by the University of Sydney in 2012 and received the Distinguished Achievement Award of the Australasian Society for Neuroscience in 2018.

Professor George Paxinos, AO (BA, MA, PhD, DSc) completed his BA at The University of California at Berkeley, his PhD at McGill University, and spent a postdoctoral year at Yale University. He is the author of almost 50 books on the structure of the brain of humans and experimental animals, including The Rat Brain in Stereotaxic Coordinates, now in its 7th Edition, which is ranked by Thomson ISI as one of the 50 most cited items in the Web of Science. Dr. Paxinos paved the way for future neuroscience research by being the first to produce a three-dimensional (stereotaxic) framework for placement of electrodes and injections in the brain of experimental animals, which is now used as an international standard. He was a member of the first International Consortium for Brain Mapping, a UCLA based consortium that received the top ranking and was funded by the NIMH led Human Brain Project. Dr. Paxinos has been honored with more than nine distinguished awards throughout his years of research, including: The Warner Brown Memorial Prize (University of California at Berkeley, 1968), The Walter Burfitt Prize (1992), The Award for Excellence in Publishing in Medical Science (Assoc Amer Publishers, 1999), The Ramaciotti Medal for Excellence in Biomedical Research (2001), The Alexander von Humbolt Foundation Prize (Germany 2004), and more.


MATERIALS AND METHODS

Animals. Twenty-nine adult female cats (8 intact and 21 spinal cord-transected) were used in the present experiments (Table1). Cats that had undergone complete low-thoracic spinal cord transection were maintained for 3–25 months.

Summary of postoperative training status and tissue analysis procedures

Spinal cord transection. During all surgical procedures, sodium pentobarbital (35 mg/kg, i.p.) was administered to each cat after pretreatment with atropine sulfate (0.9 mg/kg, s.c.) and acepromazine maleate (10 mg/kg, i.m.). Supplemental doses of the anesthetic were administered as needed during surgery to maintain a low level of arousal.

The spinal cords of the cats were transected completely as described previously (Roy et al., 1992). Briefly, a skin incision was made on the back to expose the vertebral processes between T10 and L1. A partial laminectomy was performed to expose the spinal cord between T11 and T13. Fine scissors were used to open the dura and to transect the bulk of the spinal cord. Fine forceps and miniature cotton balls were used to complete the transection while preserving the ventral artery. The completeness of the transection was easily verified as the spinal cord retracted at both ends, leaving a 3–5 mm space. Gelfoam was inserted between the two ends the area was thoroughly flushed with saline and the muscle and skin above the lesion site were closed with sutures. Postspinalization management of the spinal cats was performed as described previously (Roy et al., 1992). After all surgical procedures, the animals were placed in a warm incubator and allowed 1 d of recovery before they were returned to their cages.

Antibiotics were administered twice daily to each animal for 1 week after surgery. The cats were housed in spacious cages, two to four cats per cage. After spinal cord transection, the cage floors were covered with shredded newspaper, and the bladders and colons of the cats were expressed daily. Dry kibble and water were given ad libitum, and wet food was given once daily. All procedures were performed in accordance with the following guidelines: care and use of laboratory animals prepared by the Institute of Laboratory Animal Resources for the National Institutes of Health, the American Association for the Accreditation of Laboratory Animal Care, and the Chancellor's Animal Research Committee at the University of California, Los Angeles.

Implantation of electromyographic electrodes. Approximately 2 months before spinal cord transection, electromyographic (EMG) wires were implanted into selected flexor and extensor muscles of the hindlimbs of some cats (soleus, medial gastrocnemius, tibialis anterior, semitendinosus, vastus lateralis, gluteus medius, and iliopsoas) as described by Pierotti et al. (1989). EMG and video recordings during treadmill locomotion were initiated after 1 week of recovery and continued for the full duration of the study. EMG activity during stepping and standing was recorded periodically in animals that had undergone multiple training paradigms i.e., some cats were trained to stand and then to step and vice versa. All data presented were taken during the last training mode as shown in Table 1.

Hindlimb training and testing procedures. The training and testing procedures have been described in detail previously (de Leon et al., 1998a,b). Briefly, a cloth harness was fitted over the shoulders, between the forelimbs, and around the upper trunk. The forelimbs of the cats rested on a platform raised ∼2.5 cm above the training surface during all training and testing procedures. After spinalization, training of bipedal hindlimb stepping or standing was performed for 30 min/d, 5 d/week (for details, see de Leon et al., 1998a,b). Stepping ability during a locomotor test was measured by counting the number of plantar steps, i.e., the number of full weight-bearing steps executed on the soles of the paws in the two hindlimbs at a speed of 0.4 m/sec during a 45 sec trial. Full weight-bearing steps occurring on the dorsum of the paw were not included in the plantar step number. To train full weight-bearing bilateral hindlimb standing, the hindpaws were placed on the plantar surfaces, and the skin around the knee or ankle was patted lightly or pinched to elicit extensor reflexes. These stimuli were not used to maintain standing but were delivered when necessary to reinstate standing when the hindlimbs collapsed to a sitting, non-weight-bearing position. To train unilateral standing, weight bearing was allowed in only one hindlimb (trained limb) while the other limb (nontrained limb) was held above the training surface (de Leon et al., 1998b). The trainers held the tail only to provide lateral support during standing. To maintain lateral stability during unilateral standing, it was necessary to shift the weight of the hindquarters onto the trained limb. Typically, the trainers held the paw of the nontrained limb and moved it posteriorly and dorsally to near the base of the tail, resulting in the hindquarters leaning toward the weight-bearing limb. To evaluate standing ability after spinalization, weight bearing in one or both hindlimbs was initiated using the same stimuli that were used during training. After a weight-bearing posture was attained, the initiating stimuli were avoided, allowing the hindlimbs to stand until they collapsed to a sitting, non-weight-bearing position. Several bouts of standing were initiated to ensure a consistent performance.

EMG and kinematic data during stepping were recorded as described by de Leon et al. (1994). Briefly, raw EMG signals were amplified and recorded on a frequency modulation (FM) tape recorder (XR-510 TEAC Corp., Montebello, CA), and a camera and videocassette recorder (WV D5100 camera and AG1280P recorder Panasonic, Cypress, CA) were used to record the video signals. A Society of Motion Picture and TV Engineers time code generator (F30, Fast Forward Video, Irvine, CA) was used to synchronize video frames with the EMG signals recorded on FM tape. The EMG signals from each muscle during 10–45 sec bouts of stepping or standing were sampled into an Amiga computer at 2 kHz (de Leon et al., 1998a,b, 1999).

Tissue processing. Both intact (n = 4) and spinal cord-transected (n = 14) cats (except those used for immunoblot analysis) were perfused intracardially with 4% formaldehyde in 0.12 m phosphate buffer. The entire spinal cord, including the transection site, was removed, post-fixed in the fixative solution for 2 hr, and washed with 0.12 m phosphate buffer (four times, 30 min each). The cords then were transferred to 30% sucrose for 2 d, embedded on OCT-Tissue Tek (Miles, Elkhart, IN), and frozen on dry ice. Tissues were stored at −70°C until analysis.

Transverse sections (30 μm thick) of fixed spinal cords (L5–L7) were cut using a cryostat and collected free-floating in PBS. After washing in PBS, adjacent sections were processed for immunohistochemistry,in situ hybridization, or histochemical staining. The tissue sections used to compare the experimental groups were processed simultaneously. To minimize tissue damage that may occur with tissue handling, we processed free-floating sections using a netwell setup (75 μm mesh Costar, Cambridge, MA). Spinal cord sections in netwells were transferred sequentially to netwell trays containing appropriate solutions. Incubation with cRNA probes, antibodies, and ribonuclease A (Sigma, St. Louis, MO) and color reactions were performed in 24-well plates.

The lesion sites from each cat were evaluated for completeness of the transection by histological analysis of 20 μm sagittal sections (thawed on slides) with luxol blue (myelin) and cresyl violet (neurons and glia) stains (Kluver and Barrera, 1953).

Immunohistochemistry. Sections were processed for immunohistochemistry using an avidin–biotin complex (ABC)-peroxidase system (ABC Elite Vector Laboratories, Burlingame, CA) as described previously (Esclapez et al., 1994 Tillakaratne et al., 2000). Sections from spinal cord segments L5–L7 from control and spinal cats were incubated in 1:3000 or 1:4000 dilutions of K2 polyclonal antibody, which mainly recognizes GAD67 (Kaufman et al., 1991). The monoclonal antibody GAD6, which specifically recognizes GAD65, was used at a 1:100 dilution (Chan and Gottlieb, 1988 Erlander et al., 1991). To optimize cell body staining, we did not use detergents in the procedure. The sections were first rinsed in 0.02 m m potassium PBS (KPBS in m m : 16.5 K2HPO4, 3.5 KH2PO4, and 150 NaCl) for 30 min and then incubated with 3% normal serum (NS) diluted in KPBS for 1 hr (blocking step), followed by incubation with the antibody diluted in 1% NS for 16–20 hr at room temperature. Normal goat serum and normal horse serum (Vector Laboratories) were used for GAD67 and GAD65, respectively. Sections were rinsed three times, followed by two 10 min washes in KPBS. Sections then were incubated with the biotinylated secondary antibody (1:200 in 1% NS) for 1 hr. Sections were washed and incubated in avidin–biotin complex (A and B reagents at 1:100 in KPBS) for 1 hr. The sections were washed and reacted with the DAB and H2O2 (Sigma). Color development was monitored and ranged from 10 to 13 min. The time of the color development within an experiment was held constant. The sections were washed and mounted on superfrost slides (Fisher Scientific, Pittsburgh, PA), air-dried, and coverslipped with cytoseal (Stephens Scientific). Washings after primary, secondary, and ABC immunohistochemistry and color development were performed as described above.

In situ hybridization. Antisense and sense cRNA GAD67 probes were transcribed from subclones 13 and 16 of feline BamHI-linearized GAD67 cDNA using Sp6 RNA polymerase and digoxigenin-11-UTP (Roche Molecular Biochemicals, Indianapolis, IN) (Kaufman et al., 1986 Wuenschell et al., 1986 Tillakaratne et al., 2000). Digoxigenin-labeled RNA probes were partially digested with alkali to fragments 150–500 bases long, and their concentrations were measured as described previously (Esclapez et al., 1993, 1994Tillakaratne et al., 2000). Sections were hybridized with digoxigenin-labeled cRNA (0.2 ng/ml) for ∼16 hr at 50°C and washed as described previously (Esclapez et al., 1993, Tillakaratne et al., 2000). The hybrids were detected immunologically by incubation with alkaline phosphatase-conjugated digoxigenin antibody (Roche Molecular Biochemicals) followed by nitroblue tetrazolium chloride–5-bromo-4-chloro-3-indolylphosphate color development.

Immunoblots. L5 and L6 segments of unfixed spinal cords (four intact, four nontrained, and three step-trained spinal cord-transected cats whose spinal cords were transected 12 months earlier see Table 1) were cut into lengths of ∼3–4 mm and sonicated in homogenization buffer (100 mg tissue/ml Tillakaratne et al., 2000). The homogenization buffer consisted of 60 m m phosphate buffer, pH 7.4, 1 m m phenylmethylsulfonyl fluoride, and 0.5% Triton X-100. The homogenates were centrifuged for 15 min at 100,000 × g using a TL100 ultracentrifuge (Beckman Instruments, Palo Alto, CA), and the total protein in the supernatants was measured by the Bradford assay (Bio-Rad, Hercules, CA). For immunoblot analyses, proteins (30 μg) were separated by SDS-PAGE.

Ten 50-μm-thick L6 spinal cord transverse sections were pooled to make the protein homogenates for slot blot quantification. Spinal cord homogenates were loaded into the wells of a slot blot apparatus (Schleicher & Schuell, Keene, NH). After blotting onto nitrocellulose membranes, we detected GAD67 and β-tubulin proteins by enhanced chemiluminescence (ECL Amersham Biosciences, Arlington Heights, IL). We determined the dilutions of the primary and secondary antibodies that yielded consistent quantitative results. Blots were incubated overnight at 4°C with K2 (anti-GAD67 at 1:3000) or anti-β tubulin at 1:3000 (Chemicon, Temecula, CA) after blocking for 1 hr at room temperature with 10% nonfat milk in PBS containing 0.1% Tween 20 (PBST). After washing with PBST, the blots were incubated with the appropriate secondary antibody (1:2000) for 1 hr at room temperature. Antibody dilutions were made in PBST containing 5% nonfat milk. Washing between incubations consisted of three quick rinses, one 15 min rinse, and four 5 min rinses in PBST. Signals were visualized by ECL and exposure to x-ray film. The films then were scanned using an image analysis system (C-Imaging Compix Inc., Cranberry Township, PA) to determine the optical density of the protein bands. This protocol gives a linear standard curve using bacterially produced GAD67. Using total protein concentrations that were in the linear range of ECL detection (50–15 μg), we compared GAD67 levels after correcting for differences in loading and transfer by normalizing to β-tubulin in the same blots (Tillakaratne et al., 2000). It was possible to use the same blots two or three times with the same limits of immunodetection.

Microscopic analysis. The uniformity of tissue processing is a critical factor when analyzing the staining patterns and intensity between animals. Thus, primary fixation, post-fixation, and tissue processing were similar for the spinal cord sections prepared for immunohistochemistry or for in situ hybridization. The cat cerebellum was used as a positive control for immunohistochemistry and for in situ hybridization. The specificity of the GAD immunostaining was confirmed by the presence of GAD-immunoreactive neurons and punctate structures in the cerebellar cortex (Esclapez et al., 1993, 1994). Specificity also was confirmed by the lack of staining in tissue sections that had been incubated without the primary antibody or with the sense cRNA probe. Spinal cord transverse sections of the L5–L7 segments of intact and spinal cord-transected cats to be compared were run concurrently either for immunohistochemistry or forin situ hybridization. The time of color development within an experiment was held constant. Adjacent spinal cord sections processed for GAD67 and GAD65 were examined for the differential immunostaining reported for GAD67 but not for GAD65 after spinal cord transection (Tillakaratne et al., 2000). Cresyl violet and luxol blue staining of sections was used to evaluate the number and size of neurons.

Spinal cord sections of control and spinally transected cats, processed concurrently, were used for qualitative and semiquantitative analyses. Images to be compared were acquired with a Zeiss (Thornwood, NY) Axiophot microscope equipped with a Sony-3 CCD color camera under identical conditions (magnification, light levels, and other microscopic settings), and saved as tagged image file format files. The images were analyzed by C-Imaging software (Compix). Work files were customized for each measurement. Threshold values were set to discriminate the signal from the background. The identified objects were viewed as a green binary overlay displayed over the original image. Measurements included the staining intensity of GAD67 immunoreactivity in the lateral ventral horn of individual neurons in lamina IX and of GAD67 mRNA in individual neurons of medial and lateral laminas V and VI and the medial and lateral ventral horns. Regions of interest (ROIs subregions or individual neurons) for optical density measurements were outlined manually on the saved images. Pixels outside the ROIs were removed using the qualify feature. We added the edit option on the work file to erase, redraw, or individually remove obvious background spots before the final data acquisition. The zoom and roam feature was used on the image on display for detailed examination. The customized work files were saved and loaded to collect data from the saved images and the data were copied to Excel spreadsheets (Microsoft, Redmond, WA) and analyzed statistically. The cross-sectional areas of individual neurons were measured, and the optical density of staining was measured as the total object area. The total object area then was calculated with respect to the area of the ROI.

GAD67-positive neurons in medial lamina IX were captured at 250× at identical illumination. For the optical density measurements, individual neurons (a total of 120 neurons per cat) were outlined manually. The optical density of staining was measured as the total object area inside the outlined area (ROI). The mean GAD67 immunostaining in lamina IX in control and experimental animals (except in the unilaterally stand-trained cats) was calculated using 120 lamina IX neurons from each animal. The distribution and analysis of GAD67 immunostaining in unilaterally stand-trained cats were determined from 25 sequential transverse sections (30 μm thick, each taken at ∼120 μm) of the L5–L7 spinal segments processed for GAD67. We first compared the total object area in all of the GAD67 neurons in the contralateral and ipsilateral lamina IX region. For detailed analysis, individual GAD67-positive neurons in lamina IX of 25 sections were captured at 500× at identical illumination. The amount of GAD67 staining in individual neurons on the ipsilateral and contralateral sides (a total of 464 neurons) was measured as described above. The mean immunoreactivity per neuron between the trained and nontrained sides was compared usingt tests. The number of positively stained cells was divided into four groups using the labeling intensity values (strongest, 1.00–0.80 strong, 0.79–0.60 medium, 0.59–0.40 and low, <0.39). These values were used to add pseudocolors to neurons in the three-dimensional (3-D) representation of the GAD67 staining (see Fig. 5 also see below). We used the χ 2 test to determine whether the frequencies of cells in each intensity range were significantly different between the contralateral and ipsilateral sides (see Fig.5).

The amount of GAD67 mRNA levels in individual neurons in the medial and lateral areas of laminas V and VI and the ventral horn (from eight sections) were measured as described above. Differences in the mean immunoreactivity per neuron among the four groups were compared using the mixed model (see below). Positively stained cells were subdivided into two groups using the labeling intensity values (high, ≥0.3 and low, <0.3). We used the χ 2 test to determine whether the frequencies of cells in the intensity ranges were significantly different among the four groups (see Fig. 9).

3-D reconstruction of GAD67-immunoreactive neurons. Sequential (1:4 series) transverse spinal cord sections (30 μm thick) of the L5–L7 segments of the unilaterally stand-trained cats were processed for GAD67(1:3000, K2 antibody), GAD65 (1:100, GAD6), or cresyl violet staining. The spinal cord sections with GAD67 staining were analyzed at 31× magnification to capture the entire right or left half of the spinal cord. Overlapping images of these sections were used in Figure 4.

To make a 3-D reconstruction, we used 6 of the 25 spinal cord sections processed for immunohistochemistry (see above). The actual positions of the GAD67-positive neurons in each section were copied to separate transparencies. A digitizing program was used to incorporate x, y, and z coordinates of the GAD67-positive neurons. The position of these neurons and the corresponding optical density and size measurements then were used to construct a 3-D image of the GAD67 immunostaining with the 3-D-Studio Max program (Kinetix).

Statistical analyses. Computer-based resampling (“bootstrap”) software (version 4.0.2 Resampling Stats, Arlington, VA) and a mixed model were used to determine differences in group means (intact vs spinal, intact vs trained, and trained vs nontrained) as described previously (Efron and Tibshirani, 1993 de Leon et al., 1998a,b). A mixed model was used to study the differences in GAD67 staining among the experimental groups. The model used was: Yijg = αi + βg, whereYijg is the jth measurement of GAD67 staining for subject i in group g, αi is the intercept for subject i, and βg is the effect of group g (Efron and Tibshirani, 1993 Tillakaratne et al., 2000). The significance level was set at p < 0.05.


Evidence that 4S RNA is axonally transported in normal and regenerating rat sciatic nerves

Studies in regenerating goldfish optic nerves indicate that RNA may be axonally transported during optic nerve regeneration 14, 18, 19 . The present study was performed to determine if the axonal migration of RNA could be demonstrated during regeneration of the rat sciatic nerve.

Rats, which had only the left sciatic nerve crushed 10 days earlier, were injected bilaterally with [ 3 H]uridine into the spinal cord at segmental levels L5 and L6, thus labeling ventral horn cells giving rise to the sciatic nerve. Six, 14 and 20 days later rats were sacrificed by cardiac perfusion of saline followed by 10% formaldehyde. Formaldehyde-precipitable radioactivity, identified as [ 3 H]RNA, was 4–5 times greater in the regenerating sciatic nerve compared to the normal nerve and moved without impediment beyond the point of the crush into the regenerating portion of the nerve.

The axonal migration of free unincorporated labeled RNA precursors was also demonstrated, raising the possibility that the distribution of [ 3 H]RNA along the sciatic nerve might be entirely extra-axonal i.e., free [ 3 H]uridine is taken up by Schwann cells from the axon where it is incorporated into [ 3 H]RNA. This interpretation of the data would also result in the appearance of a proximodistal distribution of RNA associated radioactivity. To determine whether any sciatic nerve [ 3 H]RNA was due to axonal transport, rats which had only the left sciatic nerve crushed 10 days earlier were injected bilaterally with [ 3 H]uridine into the spinal cord. Fourteen days after injection, rats were sacrificed and radioactivity present in the nerve was confirmed as RNA by SDS polyacrylamide gel electrophoresis. Radioactivity in the various RNA species 14 days after intraspinal injection showed the following distribution: 28 + 18S RNA — normal39.3%±2.1 regenerating45.4%±1.6 4S RNA — normal43.0%±1.3 regenerating46.8%±2.7. Similar characterization of sciatic nerve RNA 1 or 3 days following the intravenous administration of [ 3 H]uridine gave the following distribution: 28 + 18S RNA — normal72.4%±3.0 regenerating75.0%±3.6 4S RNA — normal7.7%±1.3 regenerating10.7%±0.8.

The intraspinal injection of [ 3 H]uridine would label Schwann cell RNA and, in addition, any species of intra-axonal RNA, while intravenous injections would label Schwann cell RNA and not axonal RNA. If 4S RNA is in the axon, one would predict relatively more labeled 4S RNA following intraspinal injections than following intravenous injections. The data demonstrate an enrichment of 4S RNA in both normal and regenerating rat sciatic nerve following the intraspinal but not following the intravenous injection of labeled precursor. Therefore, we suggest that 4S RNA migrates axonally in both normal and regenerating sciatic nerves of rats.


Why Does Cauda Equina Syndrome Affect Everyone Differently?

new Lαrα
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Date Injured: March 2011

Post by Lαrα on Jan 25, 2014 4:53:45 GMT -8

Hey Guys..I know that when i first was given the news that I had Cauda Equina Syndrome I had no understanding of why this affected me in the way it does.
I can remember asking my physio to explain to me why some parts of my lower extremities had sensation more that others and why some muscles were weaker than others. why is my bladder, bowel and sexual function affected?

She explained a little but to be honest, not in a way that made enough sense to me. I searched for more understanding. from questioning and researching, gradually my understanding built up but i am always looking for more understanding and clarification.

So why does Cauda Equina Syndrome affect each of us differently?

Cauda Equina Syndrome commonly causes weakness and sensory loss to the lower limbs, the 'saddle area' and significantly affects the bladder, bowel and sexual function.

The symptoms and signs vary depending on the rate and extent of the compression, the size of the spinal canal, and the number of nerve roots involved. Because the sacral roots lie closest to the midline in the cauda equina, they bear the most damage.

A large central disc herniation, for example, would cause significant damage and symptoms including severe pain which is often worse at night.

The diagram below shows the "horse tail" that is often referred to in Cauda Equina Syndrome.This is in reference to spinal nerves within the vertebral canal. After the spinal cord tapers out, the spinal nerves continue as dangling nerve roots called cauda equina.

You will see on the diagram an aqua blue arrow that is pointing to a line that runs across the spinal nerves. This is representing an area that is compressed across the nerves. There is a black faint line above the arrow, this is showing the area the disc would be. It is situated in between L3/L4 vertebrae. Therefor the nerves below that are affected are those at the point of compression. As you can see several nerve roots are affected.

A factor in severity of longterm symptoms will depend on several factors. As many of us are aware, in the case of Cauda Equina Syndrome early intervention is crucial at the onset of symptoms.

Take a look at the illustration below..It shows which nerve roots affect the lower extremities. It will also explain why for many of us different muscles are affected more than others and why altered sensation differs in different areas.

For myself. my damage was at the point of L4/L5 disc so at the areas shown below on L3 are unaffected for me.
Sensation is affected everywhere except the for the 'strip' down the front of my legs where it is relevantly normal. My left leg is weaker then my right leg which will indicate that the nerves to this leg had more significant compression.

This video, although graphic, explains very clearly what happens at the point of compression and why we are affected differently. its a must watch!

Butiki
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Watch the video: Spinal Cord. Nervous System Biology class 10th (May 2022).