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Do neural axons extend around connections like this?

Do neural axons extend around connections like this?


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I was wondering if axons sometimes connect like this:

Note that I'm not referring to situations where the path of that network is curved and the 3rd / 4th neuron is just as close to the 1st neuron as the 2nd is, I'm talking about situations where the 3rd / 4th neuron is physically 2-3x as far from the first than the second neuron, or more. This is relevant to neural simulation.


What Is Axon Collateral? (with pictures)

Most neurons have multiple projections, or processes, that sprout from the cell body, or soma, called "neurites." Neurons have many processes, most of which are dendrites, but most neurons also have a single, unique process called an axon. Dendrites are the processes that receive signals from afferent projections of other cells, while axons are the projecting neurites, sending signals to other cells, even in distant areas of the brain or spinal cord. Axons bifurcate and can send a branch back toward its own cell's soma. These projections are known as "axon collaterals."

While a neuron has only one axon, it is common for this single process to bifurcate at multiple points along its length. An axon has a main projection, the terminal location, which determines where that cell "projects" to, and it has branches. One of the branches that bifurcates off the primary axon is the axon collateral, and it projects back toward the cell itself.

At the terminal of an axon there are "boutons" that form synapses with other cells, often on the dendritic processes of those receiving cells. There are also synapses that form along the length of the axon. In the cases of myelinated processes, synapses only form at the breaks in the myelin called "Nodes of Ranvier."

An axon collateral is often associated with feedback mechanisms that help a cell regulate its own firing pattern. Since this collateral will project back toward the cell that the axon is attached to, it can easily form synapses with the cell itself, or other, nearby cells. In the case of pyramidal, excitatory cells — common information-integration cells in the mammalian brain — are often near inhibitory interneurons. As an axon collateral can form synapses with these nearby inhibitory neurons, it serves as a regulation system. Excitatory cells excite nearby inhibitory cells that prevent them from firing quite as often.

Axon collaterals bifurcate from primary axon segments at unmyelinated regions. This makes sense because myelin functions as a neural wrap when an oligodendrocyte sends a process repeatedly around an axonal process of another cell. A single oligodendrocytic process, however, would have a difficult time wrapping around three segments joining at a single point, which is what happens when axon segments form branches.

Some scientists believe this type of collateral is a cellular preservation mechanism more than a mechanism tailored to encode firing patterns and regulate signaling information through neural pathways. This is because some excitatory cells are at risk for a toxicity response with too much activity. If an excitatory cell were to fire and the axon collateral stimulated the same cell again, a firing cascade would occur that self-propagates, causing a phenomenon known as "excito-toxicity."


Materials and Methods

Purification of NG2+ cells from the adult spinal cord.

We used a novel isolation protocol (Bai et al., 2013) to obtain highly purified NG2+ cells from adult (Ϩ weeks) C57BL/6J mice. After dissecting out the spinal cords, the surface blood vessels were removed and the tissue was dissociated with trypsin and EDTA for 30 min at room temperature. Digested tissue was spun at 800 × g for 5 min, washed three times, then placed on a Percoll gradient (GE Healthcare, catalog #17-0891-02). The gradient was centrifuged for 30 min at 2000 rpm. The cellular fractions were collected, washed, and resuspended in 1 ml of DMEM/F12 medium containing 10% fetal bovine serum. Cells were plated in a PLL-coated 75 cm 2 flask and grown at 37ଌ in 5% CO2 for 4 weeks, then passaged to purify. After passage, 99% of cells were NG2+, υ% expressed GFAP, and there was no detectable expression of mature oligodendrocyte markers, such as CNP or MBP. Most cells (84%) expressed PDGFα receptor, a marker often associated with oligodendrocyte precursor cells. The majority of cells (99%) expressed oligodendrocyte lineage markers A2B5, Olig2, and O4 (in later passages).

Culturing DRG neurons.

DRG neurons were harvested as previously described (Tom et al., 2004b). Briefly, the DRGs were removed from adult female Sprague Dawley rats (Zivic-Miller Laboratories Harlan). After the central and peripheral roots were trimmed, DRGs were incubated in a solution of collagenase II (200 U/ml Worthington Biochemicals) and dispase II (2.5 U/ml Roche) in HBSS. Digested DRGs were washed in HBSS-CMF and gently triturated three times, followed by low-speed centrifugation. The dissociated neurons were resuspended in Neurobasal A media supplemented with B-27, GlutaMAX, and penicillin–streptomycin (Invitrogen) for counting.

Longest neurite outgrowth assay.

The coverslips were coated with poly- l -lysine (PLL 0.1 mg/ml Sigma-Aldrich) overnight at room temperature, then washed with ddH20. Coverslips were bathed in laminin (5 μg/ml Invitrogen) in HBSS-CMF and incubated (37ଌ) for 2 h before plating cells. For the NG2+ cell monolayer experiments, adult mouse spinal cord NG2+ glial cells were densely plated (60,000 cells/spot) for 24 h. Cells were treated with chondroitinase ABC (ch'ase 0.1 U/ml in saline Seikagaku) for 4 h before adding dissociated DRG neurons (1500� neurons/coverslip) to the culture. DRGs, along with ch'ase in fresh medium, were added and permitted to grow for 24 h. For the laminin outgrowth coverslips, PLL-coated coverslips were bathed in 1 μg/ml or 5 μg/ml laminin in CMF and incubated at 37ଌ. After 2 h of incubation, dissociated DRG neurons were added to the culture. Outgrowth was permitted for 24 h, then the cultures were fixed with 4% paraformaldehyde and stained for NG2 (Millipore Bioscience Research Reagents) and β-III-tubulin (Sigma). For outgrowth studies, the longest neurite of each neuron growing on a complete monolayer of NG2+ cells was measured using MetaMorph software.

Entrapment assay.

The coverslips were coated with PLL and then with nitrocellulose. The coverslips were then bathed in laminin to form substrates of various concentrations [0 μg/ml, 1 μg/ml, or 5 μg/ml in Ca 2+ /Mg 2+ -free HBSS (HBSS-CMF) BTI] for 2+ h at 37ଌ. Adult mouse spinal cord NG2+ cells were plated on the coverslips at a density of 15,000 cells/coverslip. Coverslips were placed in the incubator (37ଌ). Twenty-four hours after the plating of NG2+ cells, dissociated DRG neurons were added to the culture (2000 cells/coverslip). After an additional 24 h, the cultures were fixed for 30 min with 4% paraformaldehyde, washed, and blocked in 5% natural goat serum. The fixed cells were stained for NG2, β-III-tubulin, and DAPI. Each neuron with a cell body beginning on an NG2+ cell with neurite outgrowth was imaged and quantified by counting the number of neurons capable of extending processes off the surface of the NG2+ cell. To examine the role of NG2 and other CSPGs in entrapment, chondroitinase was added in the media (0.1 U/ml) at the time of NG2+ cell plating, then again in the media at the time of adding DRG neurons. For 5 d studies, the media and ch'ase were replaced daily.

Stripe assays.

The stripe assay experiments were performed according to a protocol modified from Knöll et al., 2007. The coverslips were coated in PLL overnight at room temperature, then washed with ddH20. The coverslips were dried completely and each coverslip was placed in the center of a large Petri dish. The stripe matrix (Karlsruhe Institute of Technology, Germany) was placed on the center of the coverslip, ensuring that the entire lanes were on the coverslip. Solution A consisted of a laminin (1 μg/ml, 5 μg/ml, or 10 μg/ml Invitrogen)-aggrecan (50 μg/ml or 100 μg/ml Sigma-Aldrich) mixture or a laminin (2 μg/ml)-NG2 (7 μg/ml gift from Joel Levine, Stony Brook University, Stony Brook, NY) mixture (made with BSA conjugated with 488 in CMF). One-hundred fifty microliters of solution A was placed on the inlet channel and gently aspirated through the lanes using a flamed glass pipette, with care taken not to aspirate through the entire solution so the lanes remain covered in substrate. The coverslips were then placed in the incubator for 30 min. The solution was removed from the matrix and replaced with 2% BSA. After an additional 10 min incubation at 37ଌ, the BSA was removed from the coverslips, along with the matrix. Coverslips were placed in a 24-well plate, where they were bathed in 400 μl of solution B [low laminin (1 μg/ml) or fibronectin (5 μg/ml), with BSA in CMF] and incubated at 37ଌ for 30 min. Solution B was then removed from the wells and replaced with 2% BSA for another 10 min incubation at 37ଌ. To remove the GAG chains of the proteoglycans, chondroitinase (0.1 U/ml in saline) was added to the coverslips 4+ h before the plating of DRG neurons. Dissociated DRG neurons were then added to the coverslips at a density of 2000 cells/coverslip. Outgrowth was permitted for 48 h and then the cultures were fixed with 4% PFA. Some coverslips were immunostained for laminin (Sigma), CS56 (Sigma), and fibronectin (BD Biosciences) to ensure these components were present. All coverslips were stained with β-III-tubulin to visualize the neurons. All neurons with outgrowth twice the length of their cell bodies were imaged. Neurons with cell bodies beginning in the green lane were analyzed to see whether or not they extended processes into the black lane and vice versa.

Immunocytochemistry.

All cultures were blocked in 5% natural goat serum. All antibodies were incubated overnight at 4ଌ. The primary antibodies used included the following: anti-NG2 (Millipore), anti-GFAP (Accurate), anti-vimentin (Sigma), anti-SV2 (Developmental Studies Hybridoma Bank), anti-PSD-95 (Abcam), anti-SNAP-25 (Sigma), anti-fibronectin (BD Biosciences), anti-laminin (Sigma), DAPI (Sigma), anti-Olig2 (Millipore), anti-A2B5, and anti-O4 (generous gifts from the laboratory of Robert Miller, Case Western Reserve University, Cleveland, OH). This was followed by the appropriate secondary antibodies, Alexa Fluor 350, 488, 594, 633, or 647.

Quantification of synaptoid number in vitro between DRG axons and NG2 cells.

Pixel intensity was measured using ImageJ software for colocalization studies. Methods similar to those described by Ippolito and Eroglu, 2010 were used to quantify synapses. All images were taken with the same settings on the Zeiss LSM510 confocal microscope. An area of interest was outlined around the cell body, two times the cell body diameter in size. Using the puncta analyzer and setting the threshold for 50, the number of puncta were calculated.

Synaptic vesicle cycling assay.

Dishes were coated in poly- l -lysine overnight at room temperature, rinsed, and coated in 1 μg/ml laminin in CMF for 2 h at 37ଌ. Laminin was removed and NG2+ cells were plated densely to form a monolayer (40,000 cells in 40 μl DMEM-F12 media for 30 min at room temperature), then 3 ml of DMEM-F12 was added to the dishes and the dishes were incubated overnight. DRG neurons were added 24 h later. The dishes were incubated for 5 d, and the media was changed at day 3 to maintain the cultures. On day 5 of coculture, the media was removed and a high K + solution (5.37 m m KCl, 125 m m NaCl, 24 m m NaHCO3) containing FM dye (FM1� Invitrogen) was added to the culture for 3 min. A low Ca 2+ (0.5 m m ) solution was added to the dish, followed immediately by the addition of ADVACEP-7 (Sigma) for 5 min to quench background fluorescence. The solutions were removed and an additional wash with low Ca 2+ was performed for 10 min. The fluorescence was imaged on a Leica DMI6000 (AF) microscope. Then the solution was removed and high K + was added without dye for 3 min. The fluorescence was imaged again. ImageJ software was used to outline the entire neuron and compare the pixel intensity within the neuron before and after stimulation with or without dye.

Cell lysate for Western blot.

NG2+ cells were grown to confluency on cell culture dishes pretreated with 1× PLL for 48 h. For cells that were pretreated with ch'ase, the enzyme was added (0.1 U) to the media 1 h before adding the RIPA buffer. Cells were washed with ice-cold PBS and incubated on ice with RIPA buffer (Thermo Scientific) and 1:500 protease inhibitor cocktail (Abcam) for 1 min. Sterile cell scrapers were used to scrape cells from the cell culture dish. Cells suspended in RIPA buffer were then vortexed for 30 min at 4ଌ and spun down at 14.1 rcf for 20 min. The supernatant was saved from each group and protein concentration was assayed using a BCA kit (Thermo Scientific).

Western blot.

NG2 lysates were denatured with β-mercaptoethanol at 80ଌ for 10 min. Ten micrograms of protein was run on a 7.5% TGX gel (Bio-Rad) for 1.5 h at 125 V. Transfer occurred overnight at 12 V on PVDF membranes. PVDF membranes were blocked (Thermo Scientific Super Blocker) for 1 h shaking at room temperature and incubated at 4ଌ overnight with an NG2 antibody (NG2 Millipore). The membrane was then washed with 1× PBS and 0.1% Tween 20 three times for 15 min each at room temperature while shaking. Anti-rabbit HRP antibody (Millipore Bioscience Research Reagents) was added for2 h, shaking at room temperature. Thermo Scientific ECL was then used to develop the blot. Then the blots were washed in PBS-Tween 20 and incubated on an orbital shaker in Thermo Scientific Restore Stripping Buffer for 15 min at room temperature followed by three washes with PBS-Tween 20 for 15 min each. Blots were then blocked using Thermo Scientific Blocking Buffer for 2 h at room temperature on the orbital shaker before incubation with primary antibody anti-2B6 (Seikagaku 1:1000 dilution) overnight at 4ଌ. Before development with ECL Western blotting substrate, blots were incubated with an HRP-conjugated secondary antibody for 2 hat room temperature after three 15 min washes with PBS-Tween 20.

Cortical neuron culture and entrapment assay.

Coverslips were coated with PLL then bathed in laminin substrates of various concentrations (0 μg/ml, 1 μg/ml, or 5 μg/ml) for 2 h at 37ଌ. Adult mouse spinal cord NG2+ cells were plated on the coverslips at a density of 10,000 cells/coverslip and incubated at 37ଌ for 24 h. Whole cortices of 0- to 4-d-old BALBC/B6/129S mice (The Jackson Laboratory) were dissected in a bath of HBSS and kynurenate and incubated in a solution containing Papain, kynurenate, and cysteine for 30 min at 37ଌ. The neurons were then washed in HBSS-CMF, triturated, and filtered. After a brief low-speed centrifugation, the dissociated neurons were resuspended in Neurobasal A media containing B-27 supplement, GlutaMAX, and penicillin–streptomycin (Invitrogen) and plated on coverslips at a concentration of 2000 cells per coverslip. Coverslips were treated with an anti-mitotic agent (5𠌯luoro-2′-deoxy-uridine/uridine) at the time of neuron plating and chondroitinase ABC was added in the media (0.1 U/ml Seikagaku) where indicated. Coverslips were incubated for 24 h before fixing with 4% paraformaldehyde and immunostained.

Dorsal column crush, double-conditioning lesion, and axon labeling.

All surgeries were performed in accordance with procedures and protocols of the Animal Resource Center of Case Western Reserve University. Female adult Sprague Dawley rats (250� g) were given a dorsal column crush injury as described previously (Horn et al., 2008). Briefly, while the animals were anesthetized using inhaled isoflurane gas (2% in oxygen) and under aseptic technique, a T1 laminectomy was performed to expose the dorsal aspect of the C8 segment. The tines of Dumont #3 jeweler's forceps were inserted through a hole in the dura 1.5 mm apart and 1 mm deep. The cord was then crushed three times for 10 s each to create the lesion. Gel film was placed over the site of injury and the muscle layers were sutured with 4-0 nylon suture. Surgical staples were used to close the skin, at which time the animals received Marcaine (1.0 mg/kg) subcutaneously and buprenorphine (0.1 mg/kg) intramuscularly. Using 3000 MW Dextran-Texas Red (Invitrogen), axons were labeled through the right sciatic nerve 2 d before killing the animal. The animals were killed at 7 d, 14 d, 21 d, or 28 d (N = 6/group). The tissue was then used for either immunohistochemistry or electron microscopy. For the double-conditioning lesion studies, the dorsal column crush (DCC) procedure was performed, as mentioned above, and immediately following the injury, the right sciatic nerve was exposed and crushed. One week later, the right sciatic nerve was re-exposed and crushed a second time, 1 mm proximal to the previous nerve crush. Two weeks later (3 weeks total), the animals were labeled, perfused, and sectioned as described above.

Because NG2 staining declines over time, we conducted studies in parallel on mice expressing enhanced green fluorescent protein under an NG2-promoter. These were obtained from Dr. Dwight Bergles (Department of Neuroscience, Johns Hopkins University, Baltimore, MD). The NG2 knock-out experiments were performed on NG2 knock-out mice obtained from Dr. William Stallcup (Sanford-Burnham Medical Research Institute, La Jolla, CA). For these experiments, the DCC was performed at T12 and animals were killed after either 14 d or 28 d.

In the double-conditioning experiments, SV2 expression was compared qualitatively. Tissue from five different mice was examined for visible SV2 staining in Dextran-labeled fibers found rostral to the lesion compared with those that remained within or caudal to the lesion.

Immunohistochemistry.

The animals were perfused with 4% PFA. The cord was postfixed in 4% PFA overnight, followed by submersion in 30% sucrose overnight. The tissue was frozen in OCT mounting media and sectioned on a cryostat into 20 μm longitudinal sections. The sections were stained with anti-GFAP (Accurate Chemical and Scientific Corporation), anti-SV2 (Developmental Studies Hybridoma Bank), anti-PSD-95 (Abcam), anti-SNAP-25 (Sigma), anti-vimentin (Sigma), anti-NG2 (Millipore), or anti-fibronectin (BD Biosciences), followed by either Alexa Fluor 405, 488, or 633. Images were obtained using a Zeiss LSM510 confocal microscope.

Quantification of dystrophic endings in contact with NG2+ cells.

Tissue was used from four rats. The animals were skilled 7 d after injury as described above. Z-stack images were obtained using a Zeiss LSM510 confocal microscope. The number of Dextran-Texas Red-labeled endballs in contact with an NG2+ cell were counted and compared with the total number of endballs observed.

Electron microscopy.

For TEM analysis, cells were seeded onto a sterilized ACLAR Embedding Film (Electron Microscopy Sciences), which was cut into pieces of appropriate size to fit a 24-well plate. Adult NG2+ cells were plated on 1 μg/ml laminin at a concentration of 40,000 cells per coverslip. Twenty-four hours later DRGs (3500 per slip) were added to the coverslip and cultures were placed in a 37ଌ, 10% CO2 incubator and allowed to reach �% confluence. After the cells had reached this level of confluence (either 3 d or 5 d in coculture), the ACLAR sheets with their attached cells were immersed in fixative. The initial fixative was 2.5% glutaraldehyde in 0.1 m cacodylate buffer, pH 7.3. The specimen was postfixed in ferrocyanide-reduced 1% osmium tetroxide (Karnovsky, 1971). After a soak in acidified uranyl acetate (Tandler, 1990), the specimen was dehydrated in ethanol, passed through propylene oxide, and embedded in Poly/Bed. Thin sections were stained first with acidified uranyl acetate in 50% methanol (Tandler, 1990), then with the triple lead stain of Sato as modified by Hanaichi et al. (1986). These sections were examined in a JEOL 1200 EX electron microscope.

Statistical analysis.

All experiments were conducted in triplicate and repeated at least twice. The investigator was blinded before analyzing the data. Data were analyzed by the Student's t test, the two-proportion test, or two-way ANOVA with Tukey's post hoc test using Minitab 15 Software, or with the Kruskal–Wallis test followed by the Mann–Whitney U test with SPSS, as appropriate. All data are presented as mean ± SEM. Differences were considered statistically significant when the p value was π.05.


1 Answer 1

A: Not all neurons have many short dendrites and a single long axon. In fact, the majority of them do not have this shape. Cerebellar granule cells, which are the most numerous neurons in the brain (estimates of their total number average around 50 billion, which means that they constitute about 3/4 of the brain's neurons)[Ref 1,2,3], do not have this shape. They have an axon that branches out into long parallel fibers as shown in the diagram below (modified from the figure in Ref 2). The shapes of neurons vary greatly. This is because neurons have evolved their structures to suit their functions:

We can learn a lot about what a neuron does by looking at it’s morphology (i.e. shape). For example a neuron with large branching dendrites is likely integrating information from a large number of inputs, whereas a neuron that has dendrites that branch close to the soma, but don’t extend very far, is probably only integrating information from it near neighbors. Same thing with axons, projection neurons have long axons that allow them to communicate with neurons in distant brain regions, while a local interneuron will a short axon that will only branch locally, allowing it to talk to nearby cells. Some cells in the peripheral nervous system have the axons coming directly out of the dendrites, allowing them to efficiently convey information from one to the other. [Ref 4] (The figure below is also from Ref 4)

So, neurons have evolved their various shapes to do their functions. I think whether the one shape of these, as questioned in this thread, or many or all of these shapes are right or wrong for us to use to model neural networks is not answerable now. But I think we should try to study and learn from them because it is the best model we have. Remember, these neurons with their various shapes and connections are successful in creating one of the most wonderful phenomena in this universe: our conscious mind.

Richard H. Masland. Neuronal cell type. Current Biology. 2004 Vol 14 No 3: R497-R500.

Masland RH. Looking at Neurons Brown University. Neuroscience in Action: Understanding Our Brains and Nervous System.


RESULTS

Longitudinal pioneers of the fly CNS

Intersegmental longitudinal connections in the fly CNS are pioneered by four interneurons, pCC, MP1, dMP2 and vMP2 (Fig. 1A) (Jacobs and Goodman, 1989a Jacobs and Goodman, 1989b Lin et al., 1995 Hidalgo and Brand, 1997). dMP2 and vMP2 are born in the middle of each segmental ganglion (‘neuromere’), and begin to extend axons during stage 12. The dMP2 axon grows posteriorly and vMP2 grows anteriorly, projecting towards the vMP2 and dMP2 axons pioneering from the next segment. Simultaneously, pCC sends its axon anteriorly in association with vMP2, while the MP1 axon wraps around dMP2 and extends posteriorly in association with it. The encounter of pioneers from successive segments establishes the first intersegment connection early in stage 13. The pioneer functions of these four axons are largely redundant as all must be ablated to produce a severe defect in the mature nerve (Lin et al., 1995 Hidalgo and Brand, 1997). Below, we will focus on dMP2 and vMP2.

Notch is required to form longitudinal connections in the CNS

Removing Notch midway through embryogenesis, using a temperature-sensitive mutation, prevented formation of mature longitudinal axon tracts (Fig. 1B,C) (Giniger et al., 1993 Giniger, 1998). Examining embryos at early stage 13 revealed that the Notch phenotype is apparent at the earliest stages of the pioneering of these tracts. In temperature-shifted embryos, the dMP2 and vMP2 axons grew in the appropriate direction, but stalled, failing to make contact even by late stage 13 (Fig. 1D-F). Expressivity of this phenotype depends on the timing of the temperature shift (Crowner et al., 2003). We adjusted conditions to give a very modest expressivity (13±1.1% of early hemisegmental connections missing) to minimize unrelated CNS defects. Failure to establish intersegment connections is not due to transformation of the identities of vMP2 and dMP2, as shown by the direction of axon growth and by staining for the molecular marker Odd skipped (Spana and Doe, 1996) (data not shown). Stalling of longitudinal pioneers was also observed in Delta ts (Fig. 1G).

Notch is required for growth and guidance of longitudinal pioneers. (A) Four pioneer neurons establish intersegmental connections in the Drosophila CNS: pCC, MP1, dMP2 and vMP2. Two neuromeres and the intersegmental region between are shown. Green and white dashed lines indicate alignment of the schematic to the micrograph in B. Anterior is to the top. (B,C) Axon patterning in the mature embryonic nerve cord. Wild-type (WT B) or Notch ts (C) embryos were shifted to 32°C before pioneer axons extend, grown to stage 15/16, fixed and stained with mAb BP102. Arrows in C indicate gaps between adjacent segments in the mutant compare with wild type (arrowheads in B). Bracket indicates a single neuromere. (D,E,G) Axon patterning in early CNS. WT (D), Notch ts (E) or Delta ts (G) embryos were shifted to 32°C then fixed at mid-stage 13 and stained with mAb 22C10 to reveal pioneer axons. Yellow arrows indicate missing longitudinal connections in the mutants compare with wild type (arrowhead in D). White arrows highlight stalled dMP2 and vMP2 growth cones in Notch ts . (F) dMP2 and vMP2 neurons in Notch ts . A mid-stage 13 Notch ts embryo expressing mCD8-GFP in dMP2 and vMP2 under control of GAL4-15J2 was prepared as described for D,E and visualized with anti-GFP. Arrows indicate stalled MP2 growth cones. Wisps of projections from a stalled vMP2 growth cone are encircled. (H) Wild-type early stage 13 embryo bearing a lacZ reporter for Notch signaling activity [E(spl)mδ-lacZ], labeled with anti-β-galactosidase (β-gal red) and mAb22C10 (green). Arrows highlight the proximity of pioneer growth cones to cells with activated Notch signaling (lacZ + ). (I,J) Stage 13 embryos bearing the E(spl)mδ-lacZ reporter, labeled with anti-β-gal (red) and markers for interface glia, anti-Prospero (I, green) or anti-Repo (J, blue). Circles highlight clusters of β-gal-positive cells. (K) Notch ts embryos bearing the indicated transgenes were prepared as described for D,E and intersegmental connections that failed to form by mid-late stage 13 were quantified. All deviations from Notch ts are statistically significant (P<0.05 ANOVA). The thin vertical line on the Notch bar shows s.e.m. None of the transgenes produced dominant defects in a wild-type background under these conditions (<2% of hemisegments).

Notch is required for growth and guidance of longitudinal pioneers. (A) Four pioneer neurons establish intersegmental connections in the Drosophila CNS: pCC, MP1, dMP2 and vMP2. Two neuromeres and the intersegmental region between are shown. Green and white dashed lines indicate alignment of the schematic to the micrograph in B. Anterior is to the top. (B,C) Axon patterning in the mature embryonic nerve cord. Wild-type (WT B) or Notch ts (C) embryos were shifted to 32°C before pioneer axons extend, grown to stage 15/16, fixed and stained with mAb BP102. Arrows in C indicate gaps between adjacent segments in the mutant compare with wild type (arrowheads in B). Bracket indicates a single neuromere. (D,E,G) Axon patterning in early CNS. WT (D), Notch ts (E) or Delta ts (G) embryos were shifted to 32°C then fixed at mid-stage 13 and stained with mAb 22C10 to reveal pioneer axons. Yellow arrows indicate missing longitudinal connections in the mutants compare with wild type (arrowhead in D). White arrows highlight stalled dMP2 and vMP2 growth cones in Notch ts . (F) dMP2 and vMP2 neurons in Notch ts . A mid-stage 13 Notch ts embryo expressing mCD8-GFP in dMP2 and vMP2 under control of GAL4-15J2 was prepared as described for D,E and visualized with anti-GFP. Arrows indicate stalled MP2 growth cones. Wisps of projections from a stalled vMP2 growth cone are encircled. (H) Wild-type early stage 13 embryo bearing a lacZ reporter for Notch signaling activity [E(spl)mδ-lacZ], labeled with anti-β-galactosidase (β-gal red) and mAb22C10 (green). Arrows highlight the proximity of pioneer growth cones to cells with activated Notch signaling (lacZ + ). (I,J) Stage 13 embryos bearing the E(spl)mδ-lacZ reporter, labeled with anti-β-gal (red) and markers for interface glia, anti-Prospero (I, green) or anti-Repo (J, blue). Circles highlight clusters of β-gal-positive cells. (K) Notch ts embryos bearing the indicated transgenes were prepared as described for D,E and intersegmental connections that failed to form by mid-late stage 13 were quantified. All deviations from Notch ts are statistically significant (P<0.05 ANOVA). The thin vertical line on the Notch bar shows s.e.m. None of the transgenes produced dominant defects in a wild-type background under these conditions (<2% of hemisegments).

To ascertain where Notch is activated at this stage in wild type, we examined a transcriptional reporter for Notch activity, E(spl)mδ-lacZ (Cooper and Bray, 1999), and found β-galactosidase expression in a subset of the interface glia. These glia lined the pathway that the advancing longitudinal pioneer axons will follow and were present when those axons were growing (Fig. 1H) (Jacobs and Goodman, 1989a Hidalgo and Booth, 2000). The β-galactosidase + cells were identified as interface glia based on position and morphology (Fig. 1H), and by co-labeling for Prospero and Repo (Fig. 1I,J). The most prominent β-galactosidase expression is in glia within the neuromeres (Griffiths and Hidalgo, 2004), but at this stage there was also expression in interface glia between neuromeres, along the presumptive axon pathway (Fig. 1H, arrows).

Notch activation in interface glia was surprising because expression of Notch in neurons suppresses the late stage CNS axonal phenotype of Notch ts (Giniger, 1998 Le Gall et al., 2008). Remarkably, we now found we could efficiently rescue the early axon phenotype of Notch ts by expressing Notch either just in the glia [repo-GAL4 (all glia): 72% rescue htl-GAL4 (interface glia): 92% rescue] or just in the dMP2 and vMP2 pioneers (15J2-GAL4: 85% rescue P<0.01 in all cases t-test) (Fig. 1K). We verified that in wild type Notch is expressed in interface glia and in both dMP2 and vMP2 (data not shown). We also verified that Notch activity in both cell types is physiologically relevant using loss-of-function experiments (Fig. 1K): RNAi against Notch in interface glia significantly enhanced the Notch ts phenotype (2.5-fold P<0.005), and cell-autonomous reduction of functional Notch in dMP2 and vMP2 by cis-inhibition (Sprinzak et al., 2010) using expression of Delta lacking its intracellular domain (Itoh et al., 2003) enhanced the Notch ts phenotype by 92% (P<0.05). RNAi was not effective on the relevant timescale in dMP2 and vMP2 using 15J2-GAL4.

How can Notch rescue the growth of longitudinal pioneer axons both autonomously, acting in the pioneers, and non-autonomously, acting in the glia? We will first dissect the action of Notch in glia, and then turn to the Notch mechanism in pioneers.

Glial development in Notch ts

Notch becomes activated in interface glia by contact en passant with the axons of Delta-expressing commissural interneurons (Thomas and van Meyel, 2007). Interface glia were contacted by Delta + axons and displayed activated Notch signaling by early stage 13, in time to modulate growth of longitudinal pioneers (Fig. 2A). Moreover, expression in the glia of a Notch derivative lacking the ligand-binding domain could not rescue longitudinal axons (21% of intersegment connections were absent in Notch ts repo-GAL4UAS-Notch Δ EGF(10-12) versus 13% for Notch ts , consistent with previous evidence showing a mild dominant-negative effect of this transgene) (Le Gall et al., 2008).

ByM6A__&Key-Pair-Id=APKAIE5G5CRDK6RD3PGA" alt="Interface glia recruit a cap of Frazzled+ neuronal processes. The CNS of stage 13 Drosophila embryos was analyzed by immunofluorescence. Embryos in E-G and J are very early stage 13, other panels are mid-stage 13. (A) Stage 13 embryos bearing E(spl)mδ-lacZ reporter, labeled with anti-β-gal (red) and anti-Delta (green). A z-projection is shown in A. White arrowheads indicate Delta-positive axons coursing medially from lateral neurons. Anterior is to the right. AL shows a longitudinal cross-section at the position of the yellow triangles. AT shows a transverse cross-section at the position of the orange triangles. Thin white arrows indicate Delta-positive axons (green) as they contact Notch signaling-positive glia (red). (B) Wild-type (WT) or Notchts embryos were temperature-shifted, fixed and labeled with anti-Repo. Each white bracket indicates one segment. White arrow indicates midline. Wild-type and Notchts embryos each have approximately ten Repo+ interface glia per hemisegment. A few non-interface glia are also visible in this section. (C) Wild-type or Notchts embryos prepared as described for B but visualized with anti-Htl. (D) Wild-type or Notchts embryos prepared as described for B but visualized with anti-Prospero. Five to six Pros+ cells are visible in each segment in wild type and in Notchts (bracket). Only one focal plane is shown so not all Pros+ cells are visible in all segments. (E-E″″) Wild-type embryo expressing mCD8-GFP in interface glia, under the control of htl-GAL4, labeled with anti-GFP (green), anti-Frazzled (red) and mAb 22C10 (blue). E shows the overlay of all channels. E′-E″″ are higher magnification views of the boxed region. Thin white arrows indicate the growth cones of dMP2 and vMP2. Glia (GFP, green) and growth cones (mAb 22C10, blue) are shown in E′. Anti-Frazzled (red) is shown in E″. Anti-Frazzled (red) and growth cones (mAb 22C10, blue) are shown in E‴. E″″ is the overlay of all markers. Yellow arrows highlight the gap between the vMP2 growth cone and the glial cell. Note close apposition of dMP2 and vMP2 growth cones with Frazzled. (F) Three-dimensional rendering of an image stack of an embryo prepared as described for E. vMP2 growth cone (mAb 22C10, cyan) migrates through a bed of Frazzled+ processes (red) that separate it from the associated interface glia (GFP green). Image has been inverted to put glia beneath axon. (G-G″″) Wild-type embryo labeled with anti-Frazzled (green), anti-HRP (red) and mAb BP102 (blue). G shows overlay of all channels. A single optical section is shown (

Interface glia recruit a cap of Frazzled + neuronal processes. The CNS of stage 13 Drosophila embryos was analyzed by immunofluorescence. Embryos in E-G and J are very early stage 13, other panels are mid-stage 13. (A) Stage 13 embryos bearing E(spl)mδ-lacZ reporter, labeled with anti-β-gal (red) and anti-Delta (green). A z-projection is shown in A. White arrowheads indicate Delta-positive axons coursing medially from lateral neurons. Anterior is to the right. AL shows a longitudinal cross-section at the position of the yellow triangles. AT shows a transverse cross-section at the position of the orange triangles. Thin white arrows indicate Delta-positive axons (green) as they contact Notch signaling-positive glia (red). (B) Wild-type (WT) or Notch ts embryos were temperature-shifted, fixed and labeled with anti-Repo. Each white bracket indicates one segment. White arrow indicates midline. Wild-type and Notch ts embryos each have approximately ten Repo + interface glia per hemisegment. A few non-interface glia are also visible in this section. (C) Wild-type or Notch ts embryos prepared as described for B but visualized with anti-Htl. (D) Wild-type or Notch ts embryos prepared as described for B but visualized with anti-Prospero. Five to six Pros + cells are visible in each segment in wild type and in Notch ts (bracket). Only one focal plane is shown so not all Pros + cells are visible in all segments. (E-E″″) Wild-type embryo expressing mCD8-GFP in interface glia, under the control of htl-GAL4, labeled with anti-GFP (green), anti-Frazzled (red) and mAb 22C10 (blue). E shows the overlay of all channels. E′-E″″ are higher magnification views of the boxed region. Thin white arrows indicate the growth cones of dMP2 and vMP2. Glia (GFP, green) and growth cones (mAb 22C10, blue) are shown in E′. Anti-Frazzled (red) is shown in E″. Anti-Frazzled (red) and growth cones (mAb 22C10, blue) are shown in E‴. E″″ is the overlay of all markers. Yellow arrows highlight the gap between the vMP2 growth cone and the glial cell. Note close apposition of dMP2 and vMP2 growth cones with Frazzled. (F) Three-dimensional rendering of an image stack of an embryo prepared as described for E. vMP2 growth cone (mAb 22C10, cyan) migrates through a bed of Frazzled + processes (red) that separate it from the associated interface glia (GFP green). Image has been inverted to put glia beneath axon. (G-G″″) Wild-type embryo labeled with anti-Frazzled (green), anti-HRP (red) and mAb BP102 (blue). G shows overlay of all channels. A single optical section is shown (

0.7 μm). G′-G″″ show higher magnification views of the boxed region. G′ shows the overlay of channels G″-G″″ show separate channels as indicated. (H-I) An embryo expressing mCD8-GFP in interface glia (htl-GAL4), labeled with anti-GFP (green) and anti-Frazzled (red). H is a z-projection. Arrows highlight the correspondence of glial position and Frazzled. H′ is a magnified view of the region indicated in H. I shows a three-dimensional rendering of H, viewed obliquely. Frazzled is ventral to the glia (arrows). (J-J″) z-projection of an early stage 13 wild-type embryo expressing mCD8-GFP in interface glia (GAL4-605), labeled with anti-GFP (green) and anti-Frazzled (blue). Frazzled + cap is beginning to fill-in between successive segments (asterisks). J shows the overlay of channels. Arrows indicate an intersegmental region in which Frazzled is not yet detected but a glial cell is already present. J′ shows GFP signal in glia and J″ shows anti-Frazzled staining. At early stage 13 GAL4-605 is expressed primarily in interface glia later it will also be in some neurons.

Notch ts embryos had the normal number of interface glia, as assayed with anti-Repo (Fig. 2B). Moreover, the position and morphology of those glia appeared largely normal, and they expressed Htl and (where appropriate) Prospero (Fig. 2C,D). Thus, failure of overall glial development does not underlie the Notch axonal phenotype.


Neural activity promotes brain plasticity through myelin growth, study finds

Steve Fisch

Michelle Monje is senior author of a paper that found neuronal activity causes changes in myelin, cells that insulate nerve fibers and make them more efficient.The brain is a wonderfully flexible and adaptive learning tool. For decades, researchers have known that this flexibility, called plasticity, comes from selective strengthening of well-used synapses — the connections between nerve cells.

Now, researchers at the Stanford University School of Medicine have demonstrated that brain plasticity also comes from another mechanism: activity-dependent changes in the cells that insulate neural fibers and make them more efficient. These cells form a specialized type of insulation called myelin.

“Myelin plasticity is a fascinating concept that may help to explain how the brain adapts in response to experience or training,” said Michelle Monje, MD, PhD, assistant professor of neurology and neurological sciences.

The researchers’ findings are described in a paper published online April 10 in Science Express.

“The findings illustrate a form of neural plasticity based in myelin, and future work on the molecular mechanisms responsible may ultimately shed light on a broad range of neurological and psychiatric diseases,” said Monje, senior author of the paper. The lead authors of the study are Stanford postdoctoral scholar Erin Gibson, PhD, and graduate student David Purger.

Sending neural impulses quickly down a long nerve fiber requires insulation with myelin, which is formed by a cell called an oligodendrocyte that wraps itself around a neuron. Even small changes in the structure of this insulating sheath, such as changes in its thickness, can dramatically affect the speed of neural-impulse conduction. Demyelinating disorders, such as multiple sclerosis, attack these cells and degrade nerve transmission, especially over long distances.

Myelin-insulated nerve fibers make up the “white matter” of the brain, the vast tracts that connect one information-processing area of the brain to another. “If you think of the brain’s infrastructure as a city, the white matter is like the roads, highways and freeways that connect one place to another,” Monje said.

In the study, Monje and her colleagues showed that nerve activity prompts oligodendrocyte precursor cell proliferation and differentiation into myelin-forming oligodendrocytes. Neuronal activity also causes an increase in the thickness of the myelin sheaths within the active neural circuit, making signal transmission along the neural fiber more efficient. It’s much like a system for improving traffic flow along roadways that are heavily used, Monje said. And as with a transportation system, improving the routes that are most productive makes the whole system more efficient.

In recent years, researchers have seen clues that nerve cell activity could promote the growth of myelin insulation. There have been studies that showed a correlation between experience and myelin dynamics, and studies of isolated cells in a dish suggesting a relationship between neuronal activity and myelination. But there has been no way to show that neuronal activity directly causes myelin changes in an intact brain. “You can’t really implant an electrode in the brain to answer this question because the resulting injury changes the behavior of the cells,” Monje said.

The solution was a relatively new and radical technique called optogenetics. Scientists insert genes for a light-sensitive ion channel into a specific group of neurons. Those neurons can be made to fire when exposed to particular wavelengths of light. In the study, Monje and her colleagues used mice with light-sensitive ion channels in an area of their brains that controls movement. The scientists could then turn on and off certain movement behaviors in the mice by turning on and off the light. Because the light diffuses from a source placed at the surface of the brain down to the neurons being studied, there was no need to insert a probe directly next to the neurons, which would have created an injury.

By directly stimulating the neurons with light, the researchers were able to show it was the activation of the neurons that prompted the myelin-forming cells to respond.

Further research could reveal exactly how activity promotes oligodendrocyte-precursor-cell proliferation and maturation, as well as dynamic changes in myelin. Such a molecular understanding could help researchers develop therapeutic strategies that promote myelin repair in diseases in which myelin is degraded, such as multiple sclerosis, the leukodystrophies and spinal cord injury.

“Conversely, when growth of these cells is dysregulated, how does that contribute to disease?” Monje said. One particular area of interest for her is a childhood brain cancer called diffuse intrinsic pontine glioma. The cancer, which usually strikes children between 5 and 9 years old and is inevitably fatal, occurs when the brain myelination that normally takes place as kids become more physically coordinated goes awry, and the brain cells grow out of control.

Other Stanford co-authors of the paper are Hannes Vogel, MD, professor of pathology and of pediatrics Ben Barres, MD, PhD, professor and chair of neurobiology, as well as professor of developmental biology and of neurology and neurological sciences postdoctoral scholar Bradley Zuchero, PhD graduate students Christopher Mount, Grant Lin, Lauren Wood and Gregor Bieri and undergraduate students Andrea Goldstein, Sarah Miller and Ingrid Inema.

This research was funded by the National Institutes of Health (K08NS070926 and R01EY10257) the California Institute for Regenerative Medicine Alex’s Lemonade Stand Foundation the McKenna Claire Foundation The Cure Starts Now the Lyla Nsouli Foundation the Dylan Jewett, Connor Johnson, Zoey Ganesh, Dylan Frick, Abigail Jensen, Wayland Villars and Jennifer Kranz Memorial Funds the Ludwig Foundation the Stanford Medical Scientist Training Program the Stanford Institute for Neuro-Innovation and Translational Neurosciences the Lucile Packard Foundation for Children’s Health Stanford’s Clinical and Translational Science Award (UL1RR025744) the Howard Hughes Medical Institute Fellowship of the Life Sciences Research Foundation the Bezos Family Foundation the Child Health Research Institute at Stanford and the Anne T. and Robert M. Bass Endowed Faculty Scholarship.


Notch Signaling

Ryoichiro Kageyama , . Toshiyuki Ohtsuka , in Current Topics in Developmental Biology , 2010

2.1 Hes7 oscillations by negative feedback

Somites are segmental axial structures of vertebrate embryos that give rise to vertebral column, ribs, skeletal muscles, and subcutaneous tissues. A bilateral pair of somites forms periodically at the anterior ends of the presomitic mesoderm (PSM), located at the caudal part of embryos ( Fig. 10.2A ). During this process, mesenchymal cells of the PSM are transformed into the epithelial sheet (mesenchymal–epithelial transition) at each somite border, which segments a block of somitic cells from the PSM (segmentation). A bilateral pair of somites is formed once every 2 h in mice, every 90 min in chick, and every 30 min in zebrafish. Thus, the segmentation period differs from species to species. In addition, the period becomes longer at lower temperatures ( Jiang et al., 2000 ), indicating that it is also temperature dependent. However, within each species, the period length remains precise during development, so that the number of somites is used to identify the embryonic stages. The biological clock that regulates this periodic segmentation is called the segmentation clock ( Pourquié, 2003 ) its molecular mechanisms have been analyzed intensively in zebrafish, chick, and mice ( Dequéant and Pourquié, 2008 Mara and Holley, 2007 ).

Figure 10.2 . Hes7 oscillations in the PSM during somite segmentation. (A) The anterior ends of the PSM are segmented every 2 h in mouse embryos, forming a bilateral pair of somites (asterisk). Hes7 gene transcription is initiated in the posterior end of the PSM (phase I) and is propagated into the anterior region (phase II), stopping near the anterior end (phase III). Hes7 gene transcription and Hes7 protein expression occur in a mutually exclusive manner in all three phases, indicating that Hes7 gene transcription is repressed by Hes7 protein. (B) Dynamic Hes7 expression is caused by oscillation in individual cells [indicated by a dot in (A)]. Hes7 gene transcription and Hes7 protein expression oscillate in an antiphase manner. (See Color Insert.)

In mouse embryos, both Hes1 and Hes7 are expressed dynamically in the PSM ( Bessho et al., 2001a Jouve et al., 2000 ). The expression cycle is initiated in the posterior tip of the PSM and is propagated toward the anterior, ending near the anterior boundary of the PSM, after which segmentation of one bilateral pair of somites occurs ( Fig. 10.2A ). This dynamic expression of Hes1 and Hes7 is caused by synchronized oscillation in PSM cells ( Fig. 10.2B ). Of these two genes, Hes7 is functionally more important than Hes1 for somite segmentation: somites fail to segment and thus are severely fused in the absence of Hes7 but not in the absence of Hes1 ( Bessho et al., 2001b ). Interestingly, sustained expression of Hes7 also leads to severe somite fusion, suggesting that the oscillating expression of Hes7 is the key for maintaining periodic somite segmentation ( Niwa et al., 2007 ).

What is the mechanism producing Hes7 oscillations? In the PSM, Hes7 gene transcription and translation are mutually exclusive ( Fig. 10.2A,B ), and in the absence of a functional Hes7 protein, the Hes7 gene is continuously transcribed in the PSM ( Bessho et al., 2003 ). Activation of Hes7 promoter induces synthesis of Hes7 mRNA which is then translated to generate Hes7 protein Hes7 protein levels reach a peak within about 1 h. Hes7 protein binds to the multiple N box sequences located in the Hes7 promoter, repressing its own expression ( Fig. 10.3 ). This repression leads to disappearance of Hes7 mRNA, and then the Hes7 proteins disappear within an hour by the ubiquitin–proteasome-mediated degradation. The disappearance of Hes7 protein relieves autorepression, allowing Hes7 transcription to restart. As a result, Hes7 expression oscillates with a period of about 2 h. Thus, the negative regulation of Hes7 expression by Hes7 protein forms a negative feedback loop that is critical for maintaining oscillations.

Figure 10.3 . Hes7 oscillations are regulated by negative feedback and rapid degradation of gene products. Activation of Hes7 promoter induces synthesis of both Hes7 mRNA and Hes7 protein. Hes7 protein then binds to multiple N box sequences of Hes7 promoter and represses its own expression. This repression leads to disappearance of Hes7 mRNA and Hes7 protein because they are extremely unstable. Hes7 protein is polyubiquitinated and degraded by proteasome. Disappearance of Hes7 protein relieves negative autoregulation, allowing the next round of expression. As a result, Hes7 expression oscillates in the PSM.


Olfactory ensheathing cells: Biology in neural development and regeneration

Olfactory ensheathing cells (OECs) constitute a unique population of glia that accompany and ensheath the primary olfactory axons. They are thought to be critical for spontaneous growth of olfactory axons within the developing and adult olfactory nervous system, and have recently emerged as potential candidates for cell-mediated repair of neural injuries. Here, based on the current research, we give an overview of the biology of OECs in neural development and regeneration. This review starts with a detailed description of the cellular and molecular biological properties of OECs. Their functions in olfactory neurogenesis, olfactory axonal growth and olfactory bulb formation are sequently discussed. We also describe therapeutic applications of OECs for the treatment of a variety of neural lesions, including spinal cord injury, stroke, degenerative diseases, and PNS injuries. Finally, we address issues that may foster a better understanding of OECs in neural development and regeneration.

Research highlights

▶ Olfactory ensheathing cells (OECs) constitute a unique population of glia that accompany and ensheath the primary olfactory axons. ▶ They are thought to be critical for growth of olfactory axons within the developing and adult olfactory nervous system, and have recently emerged as potential candidates for cell-mediated repair of neural injuries. ▶ In this article we review current knowledge of OEC biology, including their molecular and cellular properties, and their role in olfactory development. ▶ We also describe therapeutic applications of OECs for a variety of neural lesions. ▶ This review, we think, may foster a better understanding of OECs biology in neural development and regeneration.


How are Interactions Between Neural Cells Established and Maintained?

The human embryo is a collection of clusters of non-specific cells, which then develop into the tissues of the adult human. In particular immature, non-specific neuroblasts must be differentiated into the highly specialised cells of the nervous system, each with a unique structure, function and synaptic interactions (Whatson 2004).

The whole function of the nervous system relies on the synaptic and other interactions between neural cells being developed appropriately and maintained throughout life. This account covers the initial development of specific neural cells from immature precursor stem cells and also the methods by which the growing specialised neurons grow along the correct routes to form their interactions with other cells, both neuronal and non neuronal.

Establishing neural cells from stem cells

Progenitor or stem cells are non-specific immature cells that are able to differentiate and form specialised cells in the adult organism. There are relatively few types of neural stem cells, with the potential to differentiate into many different types of specialised neural cells, depending on specific, localised requirements. Multipotent (able to form many types of daughter cells) neural stem cells were first identified in the early 1990s (Imitola et al. 2004) and subsequent research has focussed on specific examples of these immature progenitor cells.

Neural stem cells can give rise both to different types of mature cell, as a result of asymmetric cell division, as well as identical daughter cells via simple symmetric cell replication (Gage 2000). The exact form of the adult cell is controlled by the location of progenitors during differentiation, as well as by diffusible factors acting from a more distant site, usually the final location to which the differentiated cells will migrate.

Signalling molecules influence the differentiation of neural stem cells. For instance valproic acid (VPA) actively suppresses glial differentiation, instead encouraging the development of adult neurons, via upregulation of the neurogenic basic helix-loop-helix transcription factor (Hsieh, Gage 2004).

The mesenchymal stem cell (MSC) is derived from bone marrow and able to self regenerate and differentiate into several different types of cells in vivo (Kondo et al. 2005). MSCs are able to form both neuronal and glial cells the difference relating to the presence or absence or voltage gated ion channels indicative of the potential to develop into a full neuronal cell. Furthermore the presence of specific signalling molecules in the cell milieu can influence the development of specific types of neuronal cells. One such example involves the sonic hedgehog and retinoic acid signalling molecules, which have been shown to cause MSCs to define neurons adapted for the peripheral nervous system (Kondo et al. 2005). It is believed that this guidance of development occurs via the promotion of specific transcription factors appropriate to the adult neuronal cells. The diffusible signalling molecules would switch on the transcription factors.

A growing axon has an area of active tissue on the tip, known as the growth cone (Whatson 2004). It is the interactions between the growth cone and the surrounding structures and chemicals that govern the route along which the axon grows. Chemotactic guidance is the name given to guidance derived from close physical contact between the growth cone and structures it comes into contact with. For instance, physical obstacles such as bones or cartilage may prevent the axon from continuing along a specific route. Instead the filopodia (thin membrane extensions) would literally feel around for a route that the axonal growth could take.In the case of growth of the axons of the optic nerve, the growth of retinal neurites is influenced by the presence of laminin and fibronectin. Axons interact with these extracellular matrix components differently according to their origin with some adhering more strongly to laminin, with others showing greater allegiance to fibronectin (Whatson 2004). Hence some retinal neurites cross the optic chiasm whilst others do not.

Chemotactic guidance does not, however, operate in isolation, instead being linked to the presence of chemical signalling molecules, many of which are similar to those affecting the initial stem cell differentiation.

The idea that the gradient of chemical substances secreted by a target cell might be responsible for the ability of growth cones to find their way to a target cell was initially proposed by Ramon y Cajal more than a century ago (Ming et al. 2002). These chemotropic (literally movement in response to a chemical factor) factors are present in the extracellular matrix and act to either attract or repel the growth cone, along a chemical gradient. One example is netrin-1, which causes isolated xenopus spinal neurones to turn towards the greater concentration. However too high a concentration of netrin-1 (eg 5ng/ml-1 compared to 5 (g/ml-1) appears to desensitise the neurones, which no longer exhibit turning on a gradient (Ming et al. 2002). Netrin-1 (also known as unc-6) is also required by the worm C-elegans in order for differentiating neuroblasts to migrate along the dorsoventral axis the final direction determined by interactions between the neuroblasts and specific receptors (Hatten 2002).

Crossing the midline a detailed example

In the fruit fly drosophila the genes roundabout (deficient mutant – robo), commissureless (deficient mutant – comm) and slit (deficient mutant – sli) work together to influence axonal growth across the CNS midline (Kidd, Bland & Goodman 1999). Axons may cross the midline and become commissural or remain ipsilateral as longitudinal axons. Figure 1 below illustrates the effect that removal of each of the 3 chemotropic factors has on the appearance of longitudinal and commissural axons within the CNS axon scaffold, described as follows:

Wild type (all genes thus chemotropic factors intact) shows neat crossing of the midline, resulting in 2 commissures per segment and balanced commissural and longitudinal axons.

comm – no axons cross the midline, all remain longitudinal.

sli – axons enter the midline but then continue as longitudinal axons, within the midline.

robo – axons cross and recross the midline resulting in excessive commissural axons and very few longitudinal axons.

The authors concluded that slit acted as a short range repellent of axons with respect to the midline, thus was a chemotropic factor (Kidd, Bland & Goodman 1999). Subsequent research has shown that robo is actually repelled from the midline in response to slit, which has been found to be a large protein expressed by midline glia (Kraut, Zinn 2004).

Neuronal and Glial cell interactions

Much recent research into the methods by which developing neurons form interactions with other neural cells as well as the glial support cells, has come from study of insects such as drosophila.

Glia are crucial in the correct development of the developing eye disc in insects. The diffusible factors decapentaplegic and hedgehog both strongly influence glial proliferation, with decapentaplegic also having a role in glial motility. Glia motility is particularly crucial as axons will extend and grow in the correct direction in response to diffusible factors but will not enter the optic stalk without the chemotactic guidance of glia (Oland, Tolbert 2003).

Maintaining the connections in the adult nervous system

It has recently become apparent that, as a neuron matures, it’s responsiveness to signalling molecules such as netrin-1 changes (Salinas 2003). There is logic to this, as a maturing axon would expect to receive specific guidance cues to establish connections with other cells whereas a mature axon would not seek to continue to migrate towards the cells, rather wishing to maintain accurate connections. As there is a need to replace dead cells throughout life it would not be appropriate to completely switch off signalling molecules such as netrin-1, as every new axon would need to migrate towards the correct connections, but would then benefit from becoming effectively immune to the signalling capabilities of netrin-1 as there would be no need for further migration.

The importance of accurate migration from the stem cell germinal site to the area of axon-target interaction relates to the need for accurate synaptic connections in the adult nervous system. Organisms with a more simplistic nervous system have a low extent of migration, whilst the vertebrate nervous system involves a high degree of migration (Hatten 2002). Thus the eventual nervous system depends heavily on the accuracy of the guidance provided by both the signalling molecules and the chemotactic scaffold.

Whilst there is immense potential in understanding how stem cells develop, in terms of replicating this in order to replace damaged cells in the human adult thus far the research underlying the regulation of endogenous stem cells is in its infancy and mechanisms are poorly understood. Further, the method by which differentiated cells are able to accurately locate their target connections is also an area of current mystery.


Materials and methods

Zebrafish lines

Embryos and larvae were obtained by natural spawning from wild-type, Tg(fox D3:GFP) [12, 30], Tg(flh:eGFP) Tg(foxD3:GFP) [12], Tg(h2afz-GFP) [31], or Tg(ET16:GFP) fish (a gift from Dr Vladimir Korzh). The ET16 enhancer trap line carries a Tol2-GFP insertion and labels a subset of habenular neurons [32, 33]. Embryos were reared and staged according to standard procedures [34] and occasionally 0.002% phenylthiourea was added to the fish water from 24 hpf to inhibit pigment formation.

Dye labeling

Carbocyanine dye labeling of habenular efferent axons was performed as described previously [16].

Laser ablation

Laser ablation of parapineal precursors was performed at 24–28 hpf in Tg(flh:eGFP) Tg(foxD3:GFP) transgenic embryos as described previously [12]. Larvae were subsequently examined by laser-scanning confocal microscopy at 3 or 4 dpf to determine if any parapineal cells remained. Larvae lacking all parapineal cells were classed as 'ablated' whereas those retaining one or more parapineal cell(s) were classed as 'failed ablated'.

Focal electroporation

The electroporation technique was adapted from [35] to enable the efficient transfer of DNA to single cells or small group of cells in embryonic zebrafish CNS. Embryos at 48–72 hpf were mounted in 2% low melting point agarose (Sigma-Aldrich, St Louis, MO, USA) and using a microsurgical blade, a small chamber of agarose was cut out to expose the dorsal diencephalons/mesencephalon. Micropipettes with a tip diameter of 1–2 μm were pulled on a P-87 micropipette puller (Sutter Instrument Company, CA, USA) using AlSi glass capillaries containing a filament. Micropipettes were filled with a solution containing purified plasmid DNA resuspended in H2O at a concentration of 1 μg/μl. For most habenular neuron electroporations we used pCS2-GAP43-GFP (a gift from Dr E Amaya). GFP synthesized from this construct is localized to the cell membrane by virtue of two amino-terminal palmitoylation signals from the GAP43 protein. To visualize presynaptic terminals, we used a 1:1 mixture of pCS2-GAL4 plasmid DNA (a gift from Dr Masahiko Hibi) and pCS2-Syp:GFP-DSR [17]. This latter construct encodes both cytoplasmic DsRed fluorescent protein and a Syp-GFP fusion protein, driven from separate UAS elements. For IPN electroporations we used pCS2-lyn-Cherry, which encodes a membrane-targetted Cherry fluorescent protein (a kind gift from Henry Roehl). Micropipettes were guided into either the L or R habenula or the IPN using an MX3000 Huxley-style micromanipulator (Soma Scientific Instruments, Irvine, CA, USA) under ×40 water-immersion DIC optics (Axioskop 2 FS microscope, Carl Zeiss). The following stimulation parameters were used: 1–2 s long trains of 2 ms square pulses at 200 Hz and a potential difference of 30 V. Trains were delivered 3–5 times with approximately 0.5 s interval between trains. Pulses were generated with a Grass SD9 stimulator (Grass-Telefactor, West Warwick, RI, USA). After electroporation, embryos were cut out from the agarose and returned to embryo medium.

Whole mount in situ hybridization and immunostaining

In situ hybridization, antibody staining and histological sectioning were performed according to standard methods [36]. For antibody stainings, mouse anti-acetylated tubulin (T6793 Sigma) and rabbit anti-GFP (TP401 Torrey Pines Biolabs, San Diego, CA, USA) were used at 1:1,000 dilutions and rabbit anti-DsRed (632496 ClonTech, Palo Alta, CA, USA) was used at 1:600.

Microscopy and image manipulation

Fluorescent labeling was imaged by confocal laser-scanning microscopy (Leica SP2) using ×40 and ×63 water-immersion objective lenses. z-stacks were typically acquired at 1–2 μm intervals for epithalamic labeling and fluorescent dye-labeling of habenular axons or 0.5–1 μm intervals for imaging axonal arbors or IPN neurons labeled by electroporation. In some cases, z-stacks were deconvolved using Huygen's Essential software (Scientific Volume Imaging, Hilversum, The Netherlands). Three-dimensional projections were generated from the stack of images using Volocity software (Improvision, Coventry, UK).

In situ hybridization staining and plastic sections were photographed using a Jentopix C14 digital camera attached to a Nikon Eclipse E1000 compound microscope. For presentation, image manipulation was performed using Photoshop CS2 (Adobe) software.

Morphometric analyses

Radial distribution of neurites

To quantify the distribution of neurite branches from the center to periphery of each terminal arbor, we developed a method similar to Sholl analysis. Three-dimensional reconstructions of each arbor were orientated such that the base of the arbor lay on a flat plane and a two-dimensional image of the reconstruction, parallel to this plane, was used for further analysis. The incoming axon was cropped where it extended beyond the maximum width and length of the arbor. Next, the image was thresholded and the convex hull method was used to define the arbor perimeter (ImageJ software, US National Institutes of Health, Bethesda, Maryland, USA Hull and Circle plug-in by A Karperien and TR Roy). Using custom-written MATLAB software (The MathWorks, Inc., Natick, MA, USA), a series of 10 equally spaced concentric shells were defined, centered upon the centroid of the convex hull (see Figure 3g, for example). The number of pixels (representing axon signal) in each shell was taken as a measure of axon density. This generated a plot of cumulative fraction of axon density versus radius, for each arbor. This method is resistant to differences in the absolute area covered by the arbor and the total axonal length. Because we analyzed two-dimensional images, our method will underestimate axon density where axon segments are aligned above or below one another. This occurs rarely for L-typical arbors but is more common at the perimeter of R-typical arbors. Thus, although our method detects a greater peripheral localization of axonal length in R-typical arbors, if anything this difference between the arbor sub-types is likely to have been underestimated.

To describe the distribution profiles for the different sub-types of arbor (L-typical, R-typical and Ab-L, Ab-R) non-linear regression was used to fit fourth order polynomial models to the raw data with the y-intersect constrained to zero (at 0% radius the cumulative fraction of axon density must be zero). To compare the curves for the different arbor sub-types, we used the AIC method [37, 38]. Briefly, we used the AIC method to compare two models an AICc score is computed for a 'global' model that treats all the data from two arbor sub-types as a single data set and for a second model with individual curves fit to each data set. A large difference in the AICc scores, ΔAICc, indicates there is a high probability of the model with the lower AICc score being correct. If this is the model with separate fits for the two arbor sub-types it follows that the sub-types can be considered distinct. In the Results text we report ΔAICc and the probability that the 'individual' model, with separate polynomial fits for the two arbor sub-types, is correct.

Width/length ratio

The maximum length (measured along the AP axis) and maximum width (measured perpendicular to the AP and DV axes) were measured (Volocity, Improvision). Width/Length ratios were compared using one-way ANOVA with Tukey's post-tests for pair-wise comparisons of arbor sub-types.

Depth

The depth over which each axon elaborated its terminal arbor was measured in YZ projections made using Volocity software. For L-typical arbors located in the dIPN, depth was measured parallel to the DV axis of the brain. Because the neuropil domain of the vIPN is inclined relative to the DV axis, accurate depth measurements for ventrally located R-typical, Ab-L and Ab-R arbors were made perpendicular to the plane of the vIPN neuropil domain. Depths were compared using one-way ANOVA with Tukey's post-tests for pair-wise comparisons of arbor sub-types.

Branching

The number of branch points was counted by hand in three-dimensional reconstructions of axonal arbors. Branch points giving rise to small filopodial extensions (less than 5 μm in length) were excluded from the analysis. Average numbers of branch points were compared by one-way ANOVA with Tukey's post-tests for pair-wise comparisons of arbor sub-types.

Statistics

All statistical comparisons, nonlinear regression and comparison of curves using the AIC method were performed using Prism 4 (GraphPad Software Inc., San Diego, CA, USA).


Watch the video: Πώς μπορούμε να αναπτύξουμε νέους νευρώνες στον εγκέφαλο. TED (June 2022).


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