Short Summary Parkinson’s

Parkinson’s disease effects of physics biology chemistry and mathematics on the dynamics of the disease involved with the proper medication of direct application to the brain through teleportation of the medicineThe correct sequences of monitoring of display and calculation of effects of tomography an MRI potential effects of holographic surgeries and cephalogram free and the effect of magnetism and photonics sonar on the brain with adaptations to treat and heal the effect of previously mentioned rat model of pharmacology on human interaction with a model of regenerative biology being used on the in vitro brain with computational neuroscience and implementation of precision detail to correspond with Substantia Nigra and correlate the details involved with the afflictions control mechanisms

The details to mathematics of a linear correlation when lying in the MRI table or the operating table for 3-D Cartesian mapping math and graphing the mathematical correlation of a line drawn to represent the correlation of brain to reproductive organs with the dotted correlation representing pituitary glands pancreas and reproductive organs etcetera the line can be represented by y=(1x)=0x+sqrt(2)
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vesicle transport and axonal degeneration in CNS neurons.
Koch JC1, Bitow F1, Haack J1, d’Hedouville Z1, Zhang JN1, Tönges L1, Michel U1, Oliveira LM2, Jovin TM3, Liman J4, Tatenhorst L1, Bähr M4, Lingor P4.
Author information
Many neuropathological and experimental studies suggest that the degeneration of dopaminergic terminals and axons precedes the demise of dopaminergic neurons in the substantia nigra, which finally results in the clinical symptoms of Parkinson disease (PD). The mechanisms underlying this early axonal degeneration are, however, still poorly understood. Here, we examined the effects of overexpression of human wildtype alpha-synuclein (αSyn-WT), a protein associated with PD, and its mutant variants αSyn-A30P and -A53T on neurite morphology and functional parameters in rat primary midbrain neurons (PMN). Moreover, axonal degeneration after overexpression of αSyn-WT and -A30P was analyzed by live imaging in the rat optic nerve in vivo. We found that overexpression of αSyn-WT and of its mutants A30P and A53T impaired neurite outgrowth of PMN and affected neurite branching assessed by Sholl analysis in a variant-dependent manner. Surprisingly, the number of primary neurites per neuron was increased in neurons transfected with αSyn. Axonal vesicle transport was examined by live imaging of PMN co-transfected with EGFP-labeled synaptophysin. Overexpression of all αSyn variants significantly decreased the number of motile vesicles and decelerated vesicle transport compared with control. Macroautophagic flux in PMN was enhanced by αSyn-WT and -A53T but not by αSyn-A30P. Correspondingly, colocalization of αSyn and the autophagy marker LC3 was reduced for αSyn-A30P compared with the other αSyn variants. The number of mitochondria colocalizing with LC3 as a marker for mitophagy did not differ among the groups. In the rat optic nerve, both αSyn-WT and -A30P accelerated kinetics of acute axonal degeneration following crush lesion as analyzed by in vivo live imaging. We conclude that αSyn overexpression impairs neurite outgrowth and augments axonal degeneration, whereas axonal vesicle transport and autophagy are severely altered.
PMID: 26158517 [PubMed – in process]

Biophysical Journal
The Biophysical Society
Quantitative Imaging of Microtubule Alteration as an Early Marker of Axonal Degeneration after Ischemia in Neurons
Sotiris Psilodimitrakopoulos, Valerie Petegnief, […], and Pablo Loza-Alvarez

Additional article information

Neuronal death can be preceded by progressive dysfunction of axons. Several pathological conditions such as ischemia can disrupt the neuronal cytoskeleton. Microtubules are basic structural components of the neuronal cytoskeleton that regulate axonal transport and neuronal function. Up-to-date, high-resolution observation of microtubules in living neuronal cells is usually accomplished using fluorescent-based microscopy techniques. However, this needs exogenous fluorescence markers to produce the required contrast. This is an invasive procedure that may interfere with the microtubule dynamics. In this work, we show, for the first time to our knowledge, that by using the endogenous (label-free) contrast provided by second harmonic generation (SHG) microscopy, it is possible to identify early molecular changes occurring in the microtubules of living neurons under ischemic conditions. This is done by measuring the intensity modulation of the SHG signal as a function of the angular rotation of the incident linearly polarized excitation light (technique referred to as PSHG). Our experiments were performed in microtubules from healthy control cultured cortical neurons and were compared to those upon application of several periods of oxygen and glucose deprivation (up to 120 min) causing ischemia. After 120-min oxygen and glucose deprivation, a change in the SHG response to the polarization was measured. Then, by using a three-dimensional PSHG biophysical model, we correlated this finding with the structural changes occurring in the microtubules under oxygen and glucose deprivation. To our knowledge, this is the first study performed in living neuronal cells that is based on direct imaging of axons and that provides the means of identifying the early symptoms of ischemia. Live observation of this process might bring new insights into understanding the dynamics and the mechanisms underlying neuronal degeneration or mechanisms of protection or regeneration.

Ischemia occurs when cerebral blood flow is insufficient to meet the metabolic demand. Poor supply of oxygen (hypoxia) and glucose causes neuronal damage. Ischemia disrupts the neuronal cytoskeleton by causing changes in the phosphorylation of the microtubule-associated protein, Tau (1–5). Microtubules are main components of the neuronal cytoskeletal system. They are essential to maintain the structure of axons and dendrites and are involved in cell trafficking and axonal transport, which are crucial for neurotransmission and normal neuronal function. Under normal conditions, Tau binds to microtubules, stabilizing neuronal structure and integrity (6,7). Ischemia was shown to dephosphorylate as well to increase phosphorylation of Tau (4). A hyperphosphorylation of Tau impedes its interaction with microtubules, which are destabilized (8). Excessive phosphorylation of Tau is also assumed to be the cause of the formation of paired helical filaments-neurofibrillary tangles, seen in Alzheimer disease (7–9). In addition to changes in Tau phosphorylation, Tau can undergo proteolysis (10). Microtubule-associated protein 2 and spectrin, cytoskeleton proteins, are also degraded by activation of calpain (11). Although dephosphorylation of Tau may facilitate the binding of microtubules, there is an increase of Tau susceptibility to the protease calpain (11), and the latter may also compromise the stability of microtubules. It is likely that all these processes will also disturb the stability of the microtubules.

Monitoring structural alterations of microtubules in living neurons after exposure to ischemic conditions will contribute to better understanding of the processes leading to neuronal cell dysfunction and death. In cortical cultured neurons, a model based on oxygen and glucose deprivation (OGD) can be used to produce a controlled neuronal lesion involving progressive axon degeneration. Neuronal OGD models are now well established for in vitro investigations and are widely used in both cellular biology and preclinical stroke research (12). Identification and quantification of structural alterations induced by OGD in living primary neuron cell cultures, in the absence of external contrast, will help us to understand the dynamics of axonal degeneration and neuronal death.

A well-established high-resolution imaging technique that can be used for such a task is based on exploiting the second harmonic generation (SHG) signal originated from the axons’ microtubules (13–15). The SHG signal is produced when two excitation photons, upon interacting with matter (i.e., the sample), create one of double energy (i.e., double-frequency or at half the excitation wavelength). For this frequency doubling conversion process to occur, media-lacking inversion of symmetry is required. At the molecular level, such a condition is present in polar molecules (i.e., those possessing a permanent electric dipole) (16). Examples of such SHG active molecules are amylopectin (17), collagen triple helix (18–21), myosin (18–22), and tubulin heterodimers (23,24). Furthermore, an aligned array of such polarized molecules is able to produce, through constructive interference, an efficient SHG signal (14). This is indeed the case for starch, collagen, muscle, and microtubules, respectively. In this last particular case, SHG signal arises only from polarized microtubules such as those in the axons of neurons (13,14,25) and in other nonneuronal cell structures such as axonemes (23) and mitotic spindles (26).

This requirement on such particular molecular organization has two important implications in SHG microscopy:

Firstly, by analyzing the SHG response upon rotation of the linearly polarized excitation beam (referred to as PSHG microscopy (27,28)) it is possible to obtain structural information from the molecule itself.

Secondly, this is an intrinsic material property, not requiring any sample treatment, labeling, or staining, or genetic modification for generating contrast. This is highly advisable for in vivo microscopy studies. These properties are in contrast to other studies in ischemia where fluorescent signal was used (5).

In this study we identify, for the first time to our knowledge, early structural changes occurring in the microtubules of axons of neurons in vitro under the effect of different degrees of OGD, simulating mild or severe ischemic conditions. For that purpose, we use a three-dimensional generalization of the PSHG technique (without the use of an analyzer in the detection path) and analyze the PSHG images on a pixel-by-pixel basis. We find that in neurons exposed to a brief (60-min) period of OGD, no observable effects can be measured with PSHG when compared to the normoxic or control cultures. However, PSHG shows significant changes after exposing the neurons to 120-min OGD, which can be attributed to structural changes in microtubules. Such identification and quantification of structural neuronal changes could be used for monitoring and reflecting early signs of neuron degeneration, not only under ischemia but also in any other condition in which microtubules could be affected.

Materials and Methods
Neural cultures
Both primary cortical neuron and mixed neuron-glia cultures were prepared from 18-day-old Sprague-Dawley rat embryos (Charles River Laboratories, Wilmington, MA) as described earlier (24). Briefly, animals were anesthetized and sacrificed by cervical dislocation. All procedures were approved by the Ethical Committee for Animal Use at the University of Barcelona. Cells were resuspended in Modified Eagle’s Medium supplemented with 10% fetal calf serum and 100 μg/mL gentamycin and plated onto poly-L-lysine (5 μg/mL)-precoated glass-bottom dishes at a density of 1273 cells/mm2 for mixed cultures. Neuron-enriched cultures were prepared similarly (final density 764 cells/mm2) but medium was partly changed on the seventh day in vitro with Modified Eagle’s Medium supplemented with B27, and cytosine arabinoside was added on the fourth day in vitro to limit glial proliferation. PSHG imaging experiments in neuronal processes were performed on the seventh day in vitro in both culture types.

For performing oxygen and glucose deprivation (OGD), the culture medium of mixed neuron-glia cells was replaced by a glucose-free HEPES buffer (NaCl 135 mM, KCl 5 mM, CaCl2 2H2O 1.8 mM, MgSO4 7H2O 0.62 mM, and HEPES 10 mM). Cells were then incubated for 60 or 120 min in an hypoxic incubator (GalaxyR/RS Biotech; New Brunswick Scientific, Enfield, CT) containing 94% N2, 5% CO2, and 0,6% O2. Reoxygenation was obtained by replacing HEPES buffer with normal feeding media and cells were returned to a regular incubator (95% atmospheric air and 5% CO2). Normoxic conditions were achieved incubating the cells for 60 or 120 min with the same HEPES buffer supplemented with 5.5 mM glucose in a regular incubator (95% atmospheric air and 5% CO2). Control sister cultures, kept in a regular incubator with no medium changes, were processed in parallel. Neuron-enriched cultures were treated with 0.5 μM colchicine. Six different mixed cultures and two different neuron-enriched cultures were used. PSHG imaging experiments were performed from 2 to 4 h after reoxygenation.

For glial fibrillary acidic protein (GFAP) and Tau immunocytochemistry, control cultures were fixed in 4% paraformaldehyde for 30 min, permeabilized 5 min with Triton 0.5% in phosphate-buffered saline (PBS), blocked 30 min in PBS+3% goat serum at room temperature, and incubated overnight at 4°C with rabbit anti-GFAP 1:500 (Dako North America, Carpinteria, CA) and mouse anti-Tau 1:300 (Millipore, Billerica, MA). The following day, cultures were incubated 1 h at room temperature with anti-rabbit Alexa Fluor 546 or anti-mouse Alexa Fluor 488 secondary antibodies. Photomicrographs were acquired in a model No. IX 71 fluorescence microscope (Olympus, Melville, NY).

PSHG microscope
The setup is based on an adapted inverted microscope (TE2000-U; Nikon, Tokyo, Japan) with the laser scanning performed by a pair of galvanometric mirrors (Cambridge Technology, Nyon, Switzerland). The whole microscope unit is enclosed in a plastic box, which was heated to control the temperature at 36.7°C. A 60× (NA = 1.4, Plan Apo-Achromat; Nikon) objective was used for excitation, while for collection of the signals in the forward direction a 1.4 NA (Nikon, Japan) condenser was used. For the excitation source, we used a Kerr lens mode-locked Ti:Sapphire laser (MIRA 900fl; Coherent, Santa Clara, CA), with pulses of 160 fs at a repetition rate of 76 MHz, and operated at a central wavelength of 810 nm. After the galvanometric mirrors, we placed a linear polarizer that was followed by a zero-order λ/2 wave plate (QWPO-810; CVI Melles Griot, Leicester, UK) on a motorized rotational stage (M-060.DG; Physik Instrumente, Cranfield, Bedford, UK). This was rotated in steps to change the polarization at the sample plane. A telescope arrangement was used to ensure that a collimated beam was filling the back-aperture of the objective lens.

In the forward collection geometry, a proper mount and detection unit was implemented. This home-made mount contained a long-wave-pass dichroic beamsplitter (FF665; Semrock, Rochester, NY), a BG39 filter (Schott, Mainz, Germany), a 10-nm FWHM band pass filter centered at 405 nm (FF01-405/10-25; Semrock), and a bi-alkali photomultiplier tube (PMT, H9305-03; Hamamatsu, Hamamatsu City, Japan). In the backward collection geometry of the microscope, we choose to use only a BG39 filter (Schott) and record any signal reaching a bi-alkali PMT (H9305-05; Hamamatsu). Any effect on the depolarization of the fundamental beam introduced by the several optical components before the sample plane was assessed by measuring the mean power of the fundamental for the polarization steps of the retardation-plate, as a function of the rotation angle of an analyzer positioned parallel and perpendicular to each incoming linear polarization. The fundamental input average power, under rotation of the linear polarization (nine steps of 20°), exhibited a mean extinction coefficient ratio of 63:1 and 25:1, without and with the 60× objective, respectively.

Second harmonic generation in microtubules
As mentioned before, axons are composed of an organized array of polarized microtubules that are able to produce an efficient SHG signal. Such microtubules are composed of tubulin, which in its more basic unit consists of the alpha-beta (α-β) tubulin dimers. These heterodimers possess a permanent electric dipole (13,14). Upon interaction with high intense light, a nonlinear dipole moment μ is induced. The component μν along the molecule axis ν for a single molecule can then be described by

where Ei is the light excitation electric field component in the ν, κ, and ζ molecule coordinate system and βνκζ is the νκζ component of the microscopic third-rank hyperpolarizability tensor β. The tensor β is the one ultimately responsible for the SHG signal at the molecular level. The description of the SHG conversion is usually simplified by considering a main component βννν in one direction (16). In microtubules, βννν is associated to the α-β tubulin dimer (24). Then, the macroscopic SHG response in microtubules, which is described by a second-order susceptibility tensor χ(2), results from the coherent addition of βννν. In our approach, the susceptibility tensor element χijk in a region with a density N of α-β tubulin dimers can then be written as

where 〈⋅⋅⋅〉 denotes average; i,j,k are indices indicating the microtubule coordinate system x,y,z; and indicates a unit vector. Microtubules possesses cylindrical symmetry, resulting in only two tensor independent elements: χxxz = χyyz = χzyy =χzxx and χzzz (29), where z denotes the microtubule main symmetry axis along which it is oriented. Using an azimuth φ and a zenith θ angle to describe the ν direction and assuming a constant distribution of βννν in the azimuth plane (due to the microtubule cylindrical symmetry), the dependence of χijk with φ disappears and the ratio between the two χ(2) tensor independent elements can be expressed as (18)

where θe can be interpreted as the angle of the βννν associated to the α-β tubulin dimer with respect to the symmetry axis z of the microtubules (see Fig. 1 c).

Figure 1
Figure 1
Coordinates system of the three-dimensional PSHG biophysical model for microtubules. (a) Superposition of bright field and SHG images from a neuron with the laboratory coordinate system X, Y, Z. SHG image corresponds to microtubules. (Double headed arrow …
The PSHG technique
The induced nonlinear dipole, when excited with different incoming linear polarizations (or equivalently by rotating the sample), provides a SHG response that is characteristic of the local molecular geometrical arrangement. Specifically, we assume that the laboratory coordinate system is X-Y-Z. The laser is propagating along the laboratory Z axis and its linear polarization can be rotated in the X-Y plane in an angle α with respect to the Y axis (see Fig. 1 a). The coordinate system of the microtubules is chosen in such a way that the Y axis is contained in the X-Y plane and the Z axis coincides with the principal symmetry axis (major axis orientation) of the microtubule (Fig. 1 b). With this geometry, the relation between the lab and the microtubule frames of reference is given by the elevation angle δ and the in-plane orientation angle ϕ. Then, the detected intensity in terms of the polarization α and microtubule ϕ orientations with respect to the lab coordinate system is given by

where A is the anisotropy parameter. Note that A in Eq. 4 determines the SHG response with the polarization. To understand the nature of this differentiated SHG response, we follow the theory developed in Psilodimitrakopoulos et al. (30). Then, A can be written as

showing that there are two contributions that can cause changes in A: the elevation angle δ and the tensor ratio χzzz/χxxz.

Note that χzzz/χxxz is related to the effective orientation θe with respect to the microtubule axis z (Eq. 3), which in turn is determined by the polarization properties of the α-β tubulin dimer. A usual approach to determine the relation between the anisotropy parameter A and the angle θe is to set δ = 0 (the microtubules long axis z is assumed to lay parallel to the sample plane). Then Eq. 5 reduces to A = χzzz/χxxz, and using by Eq. 3 the effective angular orientation θe of the molecule nonlinear dipole can be found. Under this approach, any variation in A is attributed to a variation in χzzz/χzxx, and therefore to a change on the orientation of the βννν associated to the molecule, i.e., to a molecular change of the α-β tubulin dimer. Otherwise, an alternative approach can be assumed by fixing the value for χzzz/χzxx (30). This approach is equivalent to assuming that the α-β tubulin dimer does not change and any change in the experimentally retrieved anisotropy parameter A is attributed to the angle δ that describes off-plane microtubules. Then, using Eq. 5, the angle δ can be found. In a real situation, a change in the anisotropy parameter A is the result of both approaches described above, and the data must be carefully analyzed to determine the main contribution.

To retrieve the anisotropy parameter A, we use the fast-Fourier-transform (FFT)-based retrieving approach reported in Amat-Roldan et al. (31), Réfrégier et al. (32), and Gusachenko et al. (33). For that, Eq. 4 can be rewritten as a sum of Fourier components as (31–33)

where the absolute value of coefficients a0, a2, and a4 and ϕ are the unknown variables that are retrieved using the FFT-based approach. This is performed in every pixel and therefore, the result is insensible to the main orientation of the microtubule in the image. Once a0, a2, and a4 are determined, the anisotropy parameter A in microtubules can be obtained as

Note that the determination of the a0, a2, and a4 coefficients are subjected to errors such as electronic noise and others such as the effect of axial components in the focus (34), depolarization (34), and sample birefringence (35). All these errors are modeled in the error coefficient ε in Eq. 7. Nine polarizations are considered adequate to retrieve the four free parameters of Eq. 6 (22).

The above model does not differentiate between Gaussian and cone molecular distributions, it only accounts for their relevant changes. A more general model, without a priori knowledge of the symmetry and molecular distribution, has been very recently presented (36,37). Nevertheless, this is a two-dimensional model that requires the molecule axis of symmetry to lie parallel to the sample plane. Our approach consists of using a three-dimensional model, which attributes changes of the anisotropy parameter A to the tilted off-plane filaments present in thick samples.

To compare two groups we used the t-test. To compare all groups, the nonparametric Kruskal-Wallis test was performed with a post-hoc Dunn test in GraphPad Prism 4.00 for Windows (GraphPad, San Diego, CA). Differences were considered significant at P < 0.05. Results are shown as mean ± SE (standard error of the mean).

Earlier studies show that the SHG signal in neurons arises from the microtubules contained in their axons (13–15,24,25). Therefore, we sought to determine whether it was possible to detect a PSHG signal from axons in living cortical cultured neurons. Throughout our experiments, we fixed the gain in the PMT and we fixed the excitation power to 10 mW at the sample plane. In this regime, no observable damage occurred for long imaging periods of time (>2 h). We observed that the SHG signal was mostly generated in the forward direction. Although weak from a single axon, bundles of several axons offered a strong SHG signal. This was expected, because the SHG intensity is quadratically dependent upon the concentration of SHG scatterers (the microtubules), and the higher the number of axons excited within the excitation volume, the higher the generated SHG signal. This signal was lost when we treated the neurons with colchicine (a microtubule-depolymerization agent (25)), strongly indicating that the SHG was indeed microtubule-generated. Specifically, we observed ∼92% reduction in the SHG after adding colchicine in the culture (data not shown). In our hands, when neurons are cultured in the presence of glial cells, axon bundles were thicker. Therefore, we decided to perform our experiments (normoxia-OGD) in mixed cultures. The SHG was specifically recorded in axons. These cannot be mistaken for glial processes because axons are much longer and thicker than astroglial processes, as shown by GFAP-Tau immunocytochemistry in Fig. 2.

Figure 2
Figure 2
Photomicrographs of neuron and astrocyte processes in mixed cultures. Immunocytochemistry for glia fibrillary acidic protein (GFAP) (a) and Tau protein (b). Merge of A and B (c). Phase contrast image of the culture. (d) (Open arrowheads) Axons; (open …
We then proceeded to perform the PSHG analysis and retrieve the value of the anisotropy parameter, A, from cortical cultured neurons, including control (n = 9), normoxia (n = 19), and OGD applied during 60 min (n = 4) and OGD during 120 min (n = 38). Here n is the number of PSHG experiments, which are performed in four different regions in each dish. In Fig. 3 we show a typical intensity PSHG image from a control sample, acquired for each of the nine (0–160°) linear excitation polarization steps with increments of 20°. The time needed to acquire all the PSHG data was >3 min (we took three images of the same polarization, for each of the nine polarization steps). Fig. 3 shows how the SHG signal in every pixel varies with the incoming polarization. This reflects the effect of the anisotropy parameter A in the intensity modulation given by Eq. 3. In microtubules, the maximum signal is observed when the incoming linear polarization was parallel to the axons and minimum SHG signal when perpendicular. To retrieve the anisotropy parameter, the PSHG signal modulation from each pixel (the change of the SHG signal intensity due to the rotation of the excitation linear polarization observed in Fig. 3) was normalized to each pixel’s maximum intensity.

Figure 3
Figure 3
PSHG images of control cortical cultured neurons. (Arrows) Excitation linear polarization (nine steps rotation of 20°). Scale bar shows 10 μm. Each image of each polarization is composed by the average intensity of three images.
Moreover, only pixels that showed a polarization-dependent SHG signal modulation values >50% of the minimum signal were kept. The rest were considered noise and were removed. As an example, the resulting nine images of Fig. 3 were analyzed with the FFT PSHG algorithm. The result is shown in Fig. 4, a–c. The same procedure was applied to a different set of cultured neurons in which OGD was applied for 120 min. The results are displayed in Fig. 4, d–f. By simply comparing the intensity SHG images in Fig. 4, a and d, it is not possible to see any difference between the two samples. The situation is similar when the pseudocolor images of the retrieved anisotropy parameter A are compared (Fig. 4, b and e).

Figure 4
Figure 4
PSHG resulting images. (a) Noise filtered intensity image showing those pixels that are kept for analysis in control sample. (b) Image formed from the pixel-by-pixel values of the anisotropy parameter A. (c) Pixels’ histogram for the value of …
In both cases a homogeneous spatial distribution of A is observed and it is difficult to identify differences. To quantify differences between images, we can make use of image-processing techniques. A simple and useful approach is to use the image histograms, which represents the number of pixels for each tonal value in an image. In our case, the tonal value corresponds to the retrieved value of the A parameter (22). Fig. 4, c and f, corresponds to the image histogram from Fig. 4, parts b and e, respectively. In this case, the histograms are significantly different. In the case of control, the anisotropy parameter, A, has its maximum frequency at A = 1.92, having a width (defined as 2σ) of 1.88. In the case of OGD-120 min, the peak is shifted to A = 1.64, and there is a measurable increase in the histogram’s width to 1.98. This result shows that variations in the histogram of the anisotropy parameter can be used to quantify the structural differences between control and ischemic neurons by comparing their peaks and width. To compare all groups, we first pooled the normo60min and normo120min conditions because using the t-test, we found that they were not significantly different. The results obtained from control and normoxic cultures were also undistinguishable.

We thus proceed to retrieve the value of A for different neuron culture samples and analyze the histogram of each image, obtaining its peak and width. The statistical results for the two parameters are shown in Fig. 5. When comparing the results for control and normoxia samples, we found that there were no significant differences in the peak (Fig. 5 a) nor in the width (Fig. 5 b) of the anisotropy parameter A (P > 0.05 in both cases). This means that normoxia does not affect the structure of the microtubules, or else that our technique is not sensitive to any change that could have happened. The same occurs for the samples with 60 min of OGD when compared with the normoxia: there is a reduction in the peak position of A, and width is slightly reduced, but these changes are not significant (P > 0.05). The situation drastically changed for samples subjected to 120 min of OGD. Comparing the control and the 120-min OGD conditions, we found that the peak in the A shifts on average to lower values in ischemic conditions. As it can be seen in Fig. 5, in this case, the mean peak value of the anisotropy parameter A is A = 1.720. This is significantly lower than the mean peak value found for the exposure to normoxic conditions: 2.107 (P < 0.001). In addition to the clear differences in the peak values of A between normoxia and samples after 120 min of OGD, the widths of the histograms were larger in the 120-min OGD condition than in the control condition (Fig. 5 b). The difference was significant, resulting in P < 0.05.

Figure 5
Figure 5
Measurement of the anisotropy parameter A, for control, normoxia, OGD at 60 min, and OGD at 120 min. (a) The peak value of the anisotropy parameter A, Ac = 1.981 with mean ± SE = 0.079 for control, An = …
All the above results show that by either using the peak value and/or the width of the pixels’ histograms of the anisotropy parameter A, one could mark significant differences between the ischemic and normal neurons. Note that although measurements of the histograms of the different samples can be greatly affected by errors, those are systematic, and throughout these set of experiments, care was taken to ensure that they were the same for the analyzed samples. This indicates that any change in the histograms can be related to a change in the contrast mechanism in the sample, which in our case can be related to structural changes in the microtubules by considering Eq. 5. In particular, as outlined in The PSHG Technique, Eq. 5 offers two explanations on the cause that can change the histograms (30):

Case 1. The value of A can change due to a variation in the value of the tensor ratio χzzz/χzxx; and
Case 2. The histogram can change due to an increase in δ, i.e., an increase of microtubules oriented off-plane.
In the first case, a variation in the value of the tensor ratio χzzz/χzxx can be related to a change in the angle θe between the hyperpolarizability βννν orientation (associated to the α-β tubulin) and the microtubule axis z through Eq. 3. In particular, the shift in the peak distribution for the anisotropy parameter from A = 2.107–1.720 can be interpreted as an average change in the orientation from θe = 44.3–47.1°. The statistical analysis showed that this change is significant (P < 0.001).

We advance two geometrical possible interpretations of this shift: The first one is a change in the heterodimer orientation with respect to the microtubule symmetry axis z, as sketched in the change from Fig. 6, panel a, to Fig. 6, panel b. The second possibility is a compression or broadening of the heterodimer, as shown in Fig. 6 c. However, while this last possibility cannot be related to any known phenomenon affecting the heterodimer, the change of orientation is consistent with a conformational change of the β-tubulin monomer from guanosine 5-triphosphate (GTP) to guanosine 5-diphosphate (GDP) observed in nonliving samples (38–41). This conformational change favors dissociation and causes a change of 27° in the orientation of the α-β-tubulin heterodimer with respect to the microtubule symmetry axis (40). This is consistent with the average shift in the peak histograms detected after 120 min OGD, which could indicate an increase in the population of conformational changes from GTP to GDP tubulin and the start of the microtubule dissociation. In addition, these conformational changes could also be consistent with changes in the phosphorylation of the Tau microtubule-associated protein, uniquely localized in microtubules in axons—a common feature of ischemic neurons, promoting destabilization of the axon and the neuronal cytoskeleton (1–11). These evidences indicate that the PSHG analysis is able to identify microtubule degradation induced by OGD in its initial stages. It also indicates that the conformational changes observed in nonliving samples (38–41) could also occur in living neuronal cells. Importantly, these changes are not detected for 60 min OGD (mild ischemia) but appear after 120 min OGD, indicating that, after an initial resistance to ischemia, the microtubule alterations described here are early markers of axonal degeneration and neuronal death.

Figure 6
Figure 6
Biophysical interpretation of the change in the anisotropy parameter A, using Eq. 5. (a) Original α-β tubulin heterodimer, with the direction of the hyperpolarizability βννν given by the angle θ …
The second contribution, i.e., the variation in the elevation angle δ, is sketched in Fig. 6, d and e. Initially, because the samples were prepared with the axons lying mainly parallel to the focal plane of the microscope, the microtubules are expected to be oriented around δ ≈ 0. Then, Eq. 5 suggests that the value of A corresponding to the peak histogram should be close to the tensor ratio χzzz/χzxx. After this point, it is clear from Eq. 5 that an increase in the angle δ would result in a shift on the peak values of the anisotropy parameter close to A = 3. However, the shift in the histogram peak measured for 120 min OGD was in the contrary direction, i.e., to lower values of A. This is only possible if all the microtubules would decrease the value of δ, i.e., increasing the order by induction of fiber lying parallel to the focal plane—a situation that is unlikely to happen because injury tends to increase tissue disorder, at least during the initial stages. The most plausible explanation is that under OGD, the range of angles δ at which the microtubule is oriented increases, resulting in a more disordered arrangement of microtubules (transition from Fig. 6, d and e). As a result, the width of the histogram would increase (30), partially explaining the increase on the width shown in Fig. 5 b after 120 min of OGD.

In this study, we present, for the first time to our knowledge, a quantitative, label-free and all optically based technique to monitor the degeneration of microtubules in axons of living primary neuronal cells during OGD progression. This methodology allows identifying subtle structural changes of the α-β tubulin dimer (the hyperpolarizable molecule) in microtubules in axons from cultured cortical neurons. In particular, the minimally invasive, high-resolution, polarization-sensitive, second harmonic generation (PSHG) imaging was able to quantitatively assign differences between healthy and ischemic neurons.

Our experimental results and analysis showed that after 120 min of OGD, two phenomena occurred at the microtubules’ level: First, a significant change occurs in the orientation of the α-β tubulin dimer, which can be related to conformational changes from GTP to GDP tubulin and the beginning of the microtubule dissociation (38–41). Second, a significant increase in the microtubule disorder is identified. The conformational change is consistent with the well-known modification in the phosphorylation of Tau microtubule-associated protein (1–11) and with studies performed in nonliving samples (38–41). It is important to state that these changes were not observed for 60 min of OGD, showing certain neuron resistance to ischemic processes. This procedure could be used to follow not only ischemic neurons but also other types of neurodegenerative illnesses or the study of neuroprotective agents.

This work is supported by the Spanish government through the Ministry of Economy and Competitiveness, grants No. TEC2009-09698 and No. FIS2009-09928; the Ministerio de Sanidad, grant No. FIS PI081932; the Laserlab-Europe Cont. grant No. JRA4:Optobio 212025; and the Photonics for Life Networks of Excellence. This research has also been partially supported by Fundació Cellex Barcelona and partially conducted at the Institute of Photonic Sciences’ Super-Resolution Light Nanoscopy Facility.

Article information
Biophys J. 2013 Mar 5; 104(5): 968–975.
doi: 10.1016/j.bpj.2013.01.020
PMCID: PMC3870801
Sotiris Psilodimitrakopoulos,† Valerie Petegnief,‡ Nuria de Vera,‡ Oscar Hernandez,† David Artigas,†§ Anna M. Planas,‡ and Pablo Loza-Alvarez†∗
†ICFO-Institut de Ciències Fotòniques, Barcelona, Spain
‡Department of Brain Ischemia and Neurodegeneration, Institute for Biomedical Research of Barcelona, Spanish Research Council, Institut d’Investigacions Biomèdiques August Pi Sunyer, Barcelona, Spain
§Universitat Politècnica de Catalunya, Department of Signal Theory and Communications, Barcelona, Spain
Pablo Loza-Alvarez: se.ofci@azol.olbap
∗Corresponding author ; Email: se.ofci@azol.olbap
Received 2012 Jul 4; Accepted 2013 Jan 16.
Copyright © 2013 by the Biophysical Society.
This article has been cited by other articles in PMC.
Articles from Biophysical Journal are provided here courtesy of The Biophysical Society
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JCB Home > 2012 Archive > 9 January > Wang et al. 196 (1): 7
Published January 9, 2012 // JCB vol. 196 no. 1 7-18
The Rockefeller University Press, doi: 10.1083/jcb.201108111
© 2012 Wang et al.
Axon degeneration: Molecular mechanisms of a self-destruction pathway
Jack T. Wang, Zachary A. Medress, and Ben A. Barres
+ Author Affiliations
Department of Neurobiology, Stanford University School of Medicine, Stanford, CA 94305
Correspondence to Jack T. Wang:
Back to TopAbstract

Axon degeneration is a characteristic event in many neurodegenerative conditions including stroke, glaucoma, and motor neuropathies. However, the molecular pathways that regulate this process remain unclear. Axon loss in chronic neurodegenerative diseases share many morphological features with those in acute injuries, and expression of the Wallerian degeneration slow (WldS) transgene delays nerve degeneration in both events, indicating a common mechanism of axonal self-destruction in traumatic injuries and degenerative diseases. A proposed model of axon degeneration is that nerve insults lead to impaired delivery or expression of a local axonal survival factor, which results in increased intra-axonal calcium levels and calcium-dependent cytoskeletal breakdown.

As the primary signal conduit in neurons, axon fibers are on average 20,000 times larger than the cell body in length and total surface area (Friede, 1963). When the normal functions of this neuronal compartment are compromised by various insults such as trauma, blockade of axonal transport, or chemical toxicity, distinct morphological and molecular changes known as Wallerian degeneration result in cytoskeletal disassembly and granular degeneration of the axon distal to the injury site. This is followed by breakdown of the blood–brain barrier and infiltration of reactive glial cells to aid the removal of axonal and myelin debris (Fig. 1; Waller, 1850; Griffin et al., 1995; Vargas and Barres, 2007). Delaying axon degeneration prevents the progression of these subsequent glial events, suggesting that physical breakdown of the axonal cytoskeleton is required to trigger surrounding glial reactivation during Wallerian degeneration (Lunn et al., 1989; George and Griffin, 1994).

Figure 1.
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Figure 1.
Course of Wallerian axon degeneration. As early as 5–30 min after nerve injury, the axonal segments proximal (left) and distal (right) to the injury site exhibit short-distance acute axon degeneration (AAD), an event that is principally mediated by extracellular Ca2+ influx and activation of the intracellular Ca2+-dependent protease calpain. This event is followed by a slower axonal retraction and formation of axonal bulbs at the injury sites (arrowheads). For the next 24 to 48 h after injury there is a period of relative latency in which the distal axon remains morphologically stable and electrically excitable. Although beading occurs along the distal axon at irregular intervals, there are few signs of physical fragmentation. At more than 72 h after injury, rapid fragmentation and cytoskeletal breakdown occur along the full length of the distal axon, followed by increased glial (consisting primarily of astrocytes, macrophages and, in the PNS, Schwann cells) influx to clear axonal remnants (blue circles) and to possibly promote regenerative attempts by the proximal axon.

In many neurodegenerative diseases, prominent axonal pathology often precedes cell body loss in the form of “dying back,” in which axons from the synaptic regions gradually degenerate toward the cell body. Despite differences in the rate of degeneration, this process mirrors many morphological features of transected nerves, including axonal swelling, microtubule disassembly, and eventual fragmentation of the axonal cytoskeleton (Cavanagh, 1964; Griffin et al., 1995). Furthermore, molecular manipulations to delay the degeneration of severed axons also attenuate the progression of clinical pathologies in animal models of peripheral motor neuropathies, multiple sclerosis, ischemic stroke, and glaucoma (Sagot et al., 1995; Ferri et al., 2003; Samsam et al., 2003; Kaneko et al., 2006; Howell et al., 2007; Meyer zu Horste et al., 2011; Verghese et al., 2011), indicating that similar mechanisms regulate axon loss in both acute injuries and chronic diseases. Therefore, understanding the mechanisms of nerve degeneration in simple traumatic injuries allows us to model how axons are lost in more complex neurodegenerative conditions.

Much of our understanding into the nature of axon degeneration has come from imaging studies and pharmacological disruption of the process, but also from studying the Wallerian degeneration slow (WldS) mutation, which robustly delays degeneration of the distal axon from a variety of injuries including nerve transection (Lunn et al., 1989) and chemotoxic insults (Wang et al., 2001). Expression of the WldS gene product alone is sufficient to confer axon protection to wild-type neurons (Mack et al., 2001), and the ability of severed WldS axons to survive for weeks even after the cell bodies have degenerated (Deckwerth and Johnson, 1994) is evidence that axon degeneration involves mechanisms distinct from those that govern neuronal cell body death (Raff et al., 2002). This significant delay also provides an opportunity to study the mechanisms that regulate axon survival and degradation in wild-type neurons. However, distinction should be made between Wallerian axon degeneration after nerve injuries and axon pruning that occurs normally during development. Although several downstream molecules are shared, separate mechanisms control each event as WldS expression delays injury-induced axon degeneration but not axonal pruning (Hoopfer et al., 2006). The molecular pathways involved in developmental axon pruning are covered extensively in other excellent reviews (Low and Cheng, 2005; Luo and O’Leary, 2005; Saxena and Caroni, 2007). Interestingly, WldS expression does prevent the pruning of dendrites in Drosophila (Schoenmann et al., 2010; Tao and Rolls, 2011), suggesting that developmental remodeling of dendritic, but not axonal arbors involves processes akin to Wallerian degeneration. Here we focus on recent understanding of the molecular events that occur in the axon after injury, and suggest a model to explain how these events orchestrate the axonal self-destruction program.

A molecular time course of axon degeneration
Previous studies examining the sequence of events in transected axons, the simplest model of axon degeneration, reveal at least three morphologically discernible phases (Fig. 1). These consist of an acute degeneration stage that affects both the proximal and distal stumps of the axon immediately upon injury (Kerschensteiner et al., 2005), a latency period in which the distal axon remains morphologically intact and electrically excitable for brief periods (Tsao et al., 1994), and an abrupt granular degeneration phase involving the rapid fragmentation of the entire axonal cytoskeleton distal to the injury site (Lubińska, 1982; Griffin et al., 1995). These sequences of events are quickly completed by 24–48 h in vivo (Lubińska, 1977; George and Griffin, 1994) and 12–24 h in vitro in rodent axons (Glass et al., 1993; Araki et al., 2004; Wang et al., 2005). However, altering physiological conditions up to a few hours after injury, such as by lowering ambient temperature and reducing extracellular calcium levels or by genetically expressing the WldS transgene, significantly prolongs the latency period, during which the injured axons remain in a vulnerable but functional state (George et al., 1995; Tsao et al., 1999; Conforti et al., 2000). Therefore, identifying the underlying molecular events during the different morphological phases may reveal specific targets to delay or even rescue axon degeneration.

Phase I: Acute axon degeneration
It was previously thought that transected axons are structurally dormant from the time of injury until the onset of axonal fragmentation. However, recent in vivo real-time imaging studies reveal that the injured axons are far more dynamic shortly after injury. As soon as a few minutes after axotomy, the axonal segments immediately proximal and distal to the injury site rapidly degenerate by several hundred micrometers in either direction in a process that lasts between 5 and 60 min (Kerschensteiner et al., 2005). This early response to injury is followed by a slower formation of dystrophic bulb structures at the terminals of both transected ends due to accumulation of axoplasmic organelles from ongoing anterograde and retrograde transport (Fig. 1; Griffin et al., 1995).

The short, early-onset degenerative event, termed acute axonal degeneration (AAD), has been observed in both dorsal spinal sensory and optic nerves in vivo (Kerschensteiner et al., 2005; Knöferle et al., 2010). Channel-mediated influx of extracellular Ca2+ is critical for initiating AAD, as Ca2+ channel blockers prevent the early intra-axonal rise in Ca2+ and attenuate the progression of AAD. Moreover, addition of Ca2+ ionophores significantly increases the number of injured axons undergoing AAD (Knöferle et al., 2010).

The primary mechanism by which Ca2+ leads to cytoskeletal breakdown in AAD is via Ca2+-dependent activation of the serine-threonine protease calpain, which is capable of cleaving axonal neurofilament and microtubule-associated components such as spectrin and tubulin (Billger et al., 1988; Johnson et al., 1991). Increased calpain cleavage of spectrin occurs as early as 30 min after injury in vivo (Kampfl et al., 1996), corresponding to the onset of AAD. Other degradative processes such as autophagy are also triggered by the initial Ca2+ influx after axotomy (Knöferle et al., 2010) and may contribute to the severity and duration of axon loss in AAD. However, chemical inhibition of calpain fully abrogates this short distance degeneration at the severed ends of spinal cord axons (Kerschensteiner et al., 2005), indicating that calpain activity is the primary effector of AAD.

What is the purpose of AAD, and does it contribute to subsequent Wallerian degeneration of the entire distal axon? Earlier studies suggest that calpain-mediated proteolysis of neurofilaments aid in cytoskeletal restructuring and formation of growth cones in regenerating axons (Spira et al., 2003). Furthermore, the space created by this short-distance axon degeneration may enable glial proliferation at the lesion site and provide a more permissive environment for regeneration of the proximal axon. Thus, AAD may be the principal mechanism by which injured proximal axons are lost to allow neurite regrowth.

However, whether AAD affects degeneration of the distal axonal segment is less clear. Expression of the WldS transgene, which delays Wallerian degeneration, also prevents the onset of AAD (Kerschensteiner et al., 2005), suggesting that the two events may share a common mechanism. Where the events converge is not completely understood, although increased Ca2+ influx and Ca2+-dependent activation of proteases are critical in both processes (George et al., 1995; Knöferle et al., 2010). Moreover, measuring Ca2+ levels in the distal ends of transected nerves reveals distinct phases of Ca2+ elevation involving an initial Ca2+ wave front that propagates anterogradely from the injury site within seconds after axotomy (Ziv and Spira, 1993), followed by a slower rise in Ca2+ throughout the axon that occurs hours later (LoPachin et al., 1990). Although the evidence is only correlative, we surmise that such bimodal rise of Ca2+ in the axon may underlie the separate stages of axon degeneration in AAD and Wallerian degeneration. Experiments addressing whether specifically blocking AAD affects the progression of subsequent axon degeneration, such as by chelating Ca2+ or reversibly blocking Ca2+ channels only within the immediate minutes after axotomy, will help elucidate the mechanistic relationship between AAD and Wallerian degeneration.

Phase II: Latency in the distal axon
In contrast to the proximal stump, which begins to produce axoplasmic sprouts toward the lesion site only a few hours after axotomy (Kerschensteiner et al., 2005), the distal axons exhibit a period of structural quiescence after AAD, though these axons are still capable of physiological functions. For instance, severed axons of motor neurons retain their ability to conduct action potentials up to 24 h after injury in vivo, though the evoked potentials and conduction velocity progressively decay (Moldovan et al., 2009). Additionally, both anterograde and retrograde transport activities continue in the distal axon (Smith and Bisby, 1993). What is the molecular basis of this physiological latency in the injured axon, and more importantly what triggers the abrupt transition from this phase to rapid, irreversible physical degeneration? We examine the molecular events that have been shown to modulate the duration of axon survival, and discuss their roles as potential triggers for the switch to axonal degradation.

Increased intra-axonal calcium.
As an early and critical event in AAD, intra-axonal rise in Ca2+ levels is also necessary and sufficient for the subsequent Wallerian degeneration of the distal axon. Culturing neurons in a reduced calcium environment by switching to low Ca2+ media (below 200uM) or chelating external Ca2+ with EGTA robustly delays axon degeneration for 4 d after axotomy, whereas adding Ca2+ ionophores is sufficient to revert the protective phenotype and induce degeneration in uninjured neurites (Schlaepfer and Bunge, 1973; George et al., 1995). Similar to AAD, the Ca2+-dependent protease calpain is also activated in the distal axons from rising Ca2+ levels after axotomy (Glass et al., 2002); however, chemically inhibiting calpain activity only delays axon degeneration for 12–24 h in vitro (Glass et al., 1994; Wang et al., 2000). The significantly weaker axonal protection from calpain inhibition alone compared with Ca2+ chelation (Finn et al., 2000; Zhai et al., 2003) suggests that additional Ca2+-dependent proteases or pathways mediate the cytoskeletal breakdown in Wallerian degeneration.

What are the major causes of elevated intra-axonal calcium after traumatic nerve injury? As the exposed axonal membrane at the transected ends is quickly sealed by Ca2+-dependent fusion of vesicles (Eddleman et al., 1998), and as the highest axonal Ca2+ levels actually occur at significant distances away from the injury site (Ziv and Spira, 1993), the transient Ca2+ leak from openings at the cut ends is unlikely to account for the bulk of Ca2+ rise in the distal axon. Instead, the sources of axonal Ca2+ that contribute to Wallerian degeneration likely come from channel-mediated Ca2+ influx and intracellular Ca2+ release from storage sites in the injured axon itself.

Indeed, it was previously shown that L-type, but not N-type calcium channel blockers significantly delay axon degeneration for 4 d after axotomy (George et al., 1995), indicating that Ca2+ influx is required for the progression of Wallerian degeneration and is mediated by ion-specific channels. However, the signal or driving force that promotes Ca2+ entry into the injured axon is less well defined. Several reports suggest that decreased activity of Na+/K+ ATPase due to energy failure or mechanical disruption of surface membrane from nerve injuries leads to initial Na+ influx through tetrodotoxin (TTX)-sensitive Na+ channels. This then depolarizes the membrane to open voltage-gated Ca2+ channels as well as reverses the normal direction of Na+/Ca2+ exchanger activity to collectively drive Ca2+ influx (Stirling and Stys, 2010). However, addition of TTX, which blocks the inward Na+ current and attenuates intra-axonal Ca2+ levels (Wolf et al., 2001), fails to delay axon degeneration after axotomy (George et al., 1995; Press and Milbrandt, 2008), suggesting that Na+ entry is not the primary signal to trigger Ca2+ influx, or does not lead to sufficient Ca2+ entry to initiate Wallerian axon degeneration. Curiously, a decrease in intra-axonal K+ is reported to precede the rise of Ca2+ and Na+ levels in transected sciatic nerves, (LoPachin et al., 1990), raising the possibility that loss of intra-axonal K+ potential may provide a more critical electrochemical driving force for Ca2+ influx after nerve injuries.

In addition to extracellular influx, significant sources of intracellular Ca2+ are also sequestered in membranous organelles (Verkhratsky, 2005). These Ca2+ stores are tightly regulated by organelle efflux and uptake, but may be released in excess subsequent to extracellular Ca2+ entry or as a direct response to nerve injury. For instance, the mitochondria can buffer cytosolic Ca2+ through selective uniporters (Kirichok et al., 2004; Perocchi et al., 2010), and are capable of Ca2+ release by opening the permeability transition pore (PTP) complex when mitochondrial Ca2+ levels rise precipitously (Rasola and Bernardi, 2007). In fact, increasing the threshold for mitochondrial Ca2+ release by genetically ablating cyclophilin D, a critical component of the PTP in promoting mitochondrial Ca2+ efflux (Baines et al., 2005), limits white matter loss in traumatic brain injury (Büki et al., 1999) and temporarily delays axon degeneration from axotomy for up to 4 h (Barrientos et al., 2011). Moreover, at least in ischemic nerve injuries, Ca2+ is released from the ER through ryanodine and IP(3) receptors (Ouardouz et al., 2003; Nikolaeva et al., 2005), while axotomy rapidly depletes ER Ca2+ stores in the proximal axon (Rigaud et al., 2009). In more distal axons a similar system of ER-derived endomembranous tubules, collectively termed “axoplasmic reticulum” (Ellisman and Porter, 1980; Lindsey and Ellisman, 1985), expresses structural elements that functionally resemble the smooth ER in the soma (Merianda et al., 2009) and contains significant Ca2+ deposits (Henkart et al., 1978), suggesting that additional Ca2+ may be specifically stored and released from these endomembranous organelles in the axon. Further evidence of physical exchange of Ca2+ between the ER and mitochondria at regions of high Ca2+ concentration (Rizzuto et al., 1998) delineate a dynamic relationship between sites of Ca2+ storage to maintain homeostatic levels of intracellular Ca2+. Injury to the axon is likely to perturb this balance by increasing extracellular Ca2+ influx and/or triggering the release of intracellular Ca2+ stores, which overwhelms the endogenous buffering capacity and results in catastrophic rise of Ca2+ levels in the axon (Fig. 2).

Figure 2.
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Figure 2.
A molecular model of axon degeneration. (A) In the absence of injury, there is a balance between continuous somal supply via anterograde transport of an axon survival signal such as Nmnat2 and its degradation by the proteasome, resulting in sustained level of the molecule in the axon. Sufficiently high local NAD+ levels, resulting from enzymatic activity of Nmnat2, may keep axonal Ca2+ levels low by regulating the movement of Ca2+ in and out of axonal Ca2+ storage sites such as axoplasmic reticulum and mitochondria through a so-far unidentified mechanism. This regulation of Ca2+ levels in the axoplasm prevents the activation of Ca2+-dependent proteases from cleaving cytoskeletal proteins such as spectrin and preserves the structural and functional integrity of the axon. (B) Nerve injury results in impaired somal supply of Nmnat2 to the axon, resulting in diminished levels of the protein as well as axonal NAD+. Lack of NAD+-dependent regulation of Ca2+ levels lead to increased Ca2+ channel-mediated influx, reverse activity of the Na+/Ca2+ exchanger, or release of Ca2+ from internal storage sites, which together contribute to a catastrophic rise in intra-axonal Ca2+. High Ca2+ levels activate Ca2+-dependent proteases and initiate proteolytic degradation of spectrin and other axonal cytoskeletal components.

Is rising axonal Ca2+ concentration an early trigger to activate the degeneration program after injury, or an effector of more upstream signaling events? Recent evidence suggests that the latter is more likely to be the case. Events similar to Wallerian axon degeneration also occur in genetic mutants such as pmn that are impaired in axonal transport (Martin et al., 2002), where there is an absence of direct trauma and where the source of Ca2+ influx into the axon is not apparent (Coleman, 2005). Moreover, exogenous addition of Ca2+ is sufficient to abolish WldS-mediated axonal protection (Glass et al., 1994), indicating that the calcium-dependent activities in the axon are downstream of molecular components that suppress the activation of Wallerian degeneration. Thus, although Ca2+ signaling is clearly critical for axonal degradation, more upstream cellular processes likely trigger the increased intra-axonal Ca2+ levels by promoting channel-mediated influx or modulating intracellular Ca2+ buffering and release, or both, while subsequent Ca2+-dependent proteolytic events mediate the physical destruction of the axon.

Intra-axonal signaling of axon death/survival signals.
The above conclusion invites the question of what, if any, signaling event is sufficient to trigger the Ca2+-dependent degeneration program in the axon? Two potential mechanisms may be used by the cell to signal nerve injury and initiate axon degeneration. First, axonal injury may directly activate or uncage a death signal to trigger a Ca2+-dependent degeneration program. Alternatively, nerve injury may impair the transport or expression of an axonal trophic factor, causing the molecule to fall below a critical threshold to support axonal integrity and function (Lubińska, 1982; Coleman, 2005). There is recent evidence supporting both hypotheses.

Consistent with the “death signal” hypothesis, recent reports implicate a network of kinases that promotes axonal degeneration after nerve injuries. For instance, loss-of-function mutations in mammalian DLK or its Drosophila homologue wallenda, a member of the mitogen-activated kinase kinase kinase family, as well as use of a chemical inhibitor of JNK kinase, a target of DLK, all result in axon protection up to 48 h after axotomy (Miller et al., 2009). Moreover, both genetic knockdown by shRNA and chemical inhibition of GSK3 or IKKB were also sufficient to delay axon degeneration by 24 h in vitro after the same injury to neurites (Gerdts et al., 2011). In all these events, the axonal protection occurs only when the kinase activity is inhibited between time of axotomy and up to 3 h after injury (Miller et al., 2009), suggesting that these molecules may function as early sensors of axonal injury. Yet it is unclear how injury leads to activation of these kinases, and whether increased kinase activity is sufficient to induce spontaneous axon degeneration or abolish WldS-mediated axon protection. Moreover, the extent of axon protection from inhibiting these molecules alone remains considerably weaker than that from WldS expression or Ca2+ chelation, indicating that activation of these molecules may be one of several parallel injury response events rather than the principal trigger of axon degeneration. In another study supporting the presence of an axonal self-destruction signal, Nikolaev et al. (2009) observed that trophic starvation of neurons leads to cleavage and secretion of APP, a precursor of β-amyloid protein and a marker of axonal damage (Coleman, 2005). The secreted APP binds to the TNF family receptor DR6 as an autocrine signal to trigger a BAX-dependent, caspase-6–mediated breakdown of axonal cytoskeleton (Nikolaev et al., 2009). However, blocking the release of APP or chemically inhibiting caspase-6 fails to delay axon degeneration after axotomy (Vohra et al., 2010), suggesting that trophic starvation triggers a distinct axonal self-destruction program from that caused by traumatic injuries.

At the same time, there is also evidence suggesting that a constitutively transported or expressed factor normally supports axonal survival, and its absence or degradation after injury triggers axon degeneration. Recently, Gilley and Coleman (2010) observed that focal inhibition of protein translation in the cell body, but not in the axon, results in spontaneous axon degeneration of uninjured neurites. This suggests that synthesis of a protein factor in the soma and its delivery to the axon, rather than local axonal translation, maintains axon viability. Remarkably, the group identifies Nmnat2, a primarily ER and Golgi-localized protein that catalyzes the rate-limiting step of NAD+ synthesis (Berger et al., 2005), as one of the factors required for the survival of wild-type axons. In the CNS, Nmnat2 is a highly neuronal-specific protein (Cahoy et al., 2008). It is expressed in the axons and its expression quickly decreases within 4 h after axotomy or after blockade of axonal transport (Gilley and Coleman, 2010). This turnover time for the protein correlates with the latent period between axon injury and the initial appearance of axonal blebbing (Beirowski et al., 2004). Moreover, depletion of Nmnat2, but not other isoforms of Nmnat enzyme using siRNA specifically induces degeneration in uninjured axons, while overexpression of Nmnat2 delays the degeneration of wild-type axons from axotomy for up to 48 h (Gilley and Coleman, 2010; Yan et al., 2010). These exciting findings indicate that Nmnat2 is continuously required to promote endogenous axon survival, and that decreased level of the protein—caused by impaired axonal transport from the soma, local degradation in the axon, or both—leads to failure to suppress a default axon degeneration program (Fig. 2).

Ubiquitin–proteasome system.
Previous studies demonstrate that inhibiting the ubiquitin–proteasome system (UPS) by blocking proteasome activity prevents axon pruning during development (Watts et al., 2003) and delays axon degeneration from injury up to 3 d in vivo in mammalian nerves and 5–10 d in Drosophila axons (Zhai et al., 2003; MacInnis and Campenot, 2005; Hoopfer et al., 2006). How does proteasome inhibition protect axons, and through what mechanism does axonal injury affect ubiquitination or proteasome activity?

As the primary function of the proteasome is to regulate protein turnover, a likely explanation is that proteasome inhibition helps sustain intracellular levels of molecules that promote axonal survival by maintaining transport of the factor to the axon or by preventing the degradation of the molecule itself. Consistent with the first mechanism, proteasome inhibition in axotomized nerves attenuates microtubule disassembly and preserves axonal transport by preventing the turnover of microtubule-associated factors such as MAP1 and tau that help stabilize the microtubule network (Zhai et al., 2003). Moreover, inhibiting proteasome activity may also directly interfere with the degradation of a putative axonal survival factor such as Nmnat2. Indeed, the turnover of Nmnat2 is shown to be dependent on proteasome activity as its levels in the transected axon remain high when proteasome activity is blocked (Gilley and Coleman, 2010). Thus, in axonal injuries where the somal supply of the factor is lost, the level of the survival factor in the axon is then solely determined by its rate of proteasome-dependent degradation, whereas proteasome inhibition helps sustain sufficient levels of the factor in the axon to delay the onset of axon degeneration (Fig. 2).

Phase III: Granular fragmentation
The latent phase of Wallerian degeneration is followed by an abrupt switch to explosive, asynchronous fragmentation that is completed along the length of the axolemma within 1–2 h from the onset, with a rate of up to 24 mm/h in vivo and 0.4 mm/h in culture (Sievers et al., 2003; Beirowski et al., 2005). Interestingly, focal, severe injuries such as axotomy result in a proximal-to-distal direction of axon degeneration, whereas in more chronic injuries the axons degenerate from synaptic ends to the cell body in a retrograde pattern (Beirowski et al., 2005). The basis for the injury-dependent differences in the direction of degeneration is not fully understood. A recent report suggests that increased macroautophagy activity that is dependent on AKT/mTOR signaling mediates retrograde axon degeneration in dopaminergic neurons after acute chemotoxic injury (Cheng et al., 2011). However, whether this mechanism participates in Wallerian degeneration and whether the signaling events occur broadly in other CNS neurons is unclear. On the other hand, the proximal-to-distal direction of degeneration from acute nerve transection may be accounted for by the survival factor hypothesis, as depletion of the survival factor would occur at the proximal tip of the distal stump first if anterograde transport continues in the axon (Lubińska, 1982). The segment closest to the cut site would likely experience the earliest loss of the survival factor, and therefore be the first to undergo degeneration after axotomy.

Delayed axonal degeneration: Molecular mechanisms of WldS axon protection
The discovery of the WldS mouse mutant, which robustly protects both CNS and PNS nerves from physical injury, chemotoxic insult, and neurodegenerative conditions (Lunn et al., 1989; Perry et al., 1991; Wang et al., 2001; Ferri et al., 2003; Howell et al., 2007) changed our understanding of the nature of axon degeneration in injury and disease. In contrast to complete fragmentation of wild-type axons within 48 h after axotomy, the transected WldS axons remain structurally intact and electrically excitable for weeks in vivo and up to a week in culture (Lunn et al., 1989; Ludwin and Bisby, 1992; Wang et al., 2005). Transgenic expression of the WldS protein also leads to axon protection in many species, including rats (Adalbert et al., 2005) and Drosophila (Hoopfer et al., 2006; MacDonald et al., 2006). This protection, which is dose dependent (Mack et al., 2001), also extends to the synapses as WldS expression preserves synaptic integrity and function in the peripheral neuromuscular junction after axotomy (Wang et al., 2001; Adalbert et al., 2005) as well as in the CNS striatum after cortical lesions (Gillingwater et al., 2006a; Wright et al., 2010). Continued axon protection after grafting WldS nerves to wild-type hosts shows that WldS-mediated protection is intrinsic to the axon (Glass et al., 1993), and the mechanism is distinct from simply inhibiting the classical apoptotic pathways that govern cell body degeneration (Deckwerth and Johnson, 1994; Sagot et al., 1995; Finn et al., 2000). Thus, determining the molecular components of the degeneration pathway that WldS is interfering with is critical to understanding how axons are normally lost after injury.

The WldS mutation results in the formation of a chimeric gene product consisting of the first 70 amino acids of a ubiquitination factor (Ube4b) and the full sequence of a NAD+ synthetic enzyme (Nmnat1; Mack et al., 2001). The N-terminal 70 amino acids of Ube4b within WldS contain no enzymatic activity, though it contains a binding site for VCP (Laser et al., 2006), a cytoplasmic protein with diverse cellular functions (Wang et al., 2004). This VCP binding domain, along with the enzymatic activity of Nmnat1, are both required for WldS-mediated axon protection (Avery et al., 2009; Conforti et al., 2009). However, overexpression of mutant Nmnat that lacks the enzymatic domain in Drosophila still protects photoreceptor cells from SCA-1–induced degeneration (Zhai et al., 2006). This nonenzymatic protection by Nmnat has been shown to be a consequence of chaperone functions of the protein (Zhai et al., 2008) and suggests that, at least in nonmammalian species, Nmnat is able to confer neuroprotection independent of its enzymatic properties. However, whether this phenotype is axonal specific or an indirect result of broader protection of the cell bodies remains unclear.

Although the WldS protein is predominantly localized in the nucleus due to endogenous nuclear localization of Nmnat1, emerging evidence instead points to the trace amount of extranuclear WldS protein as the critical component for axon protection (Coleman and Freeman, 2010). Indeed, WldS protein has recently been detected in the axoplasm and in axonal organelles including the mitochondria and phagosomes (Beirowski et al., 2009; Yahata et al., 2009). Moreover, misexpression of Nmnat1 alone outside of the nucleus by deleting its nuclear localizing sequence (Beirowski et al., 2009; Sasaki et al., 2009), virally transducing Nmnat1 in injured axons (Sasaki and Milbrandt, 2010), and fusing it to the N-terminal sequence of APP protein to increase expression in axonal compartments (Babetto et al., 2010) all lead to robust axon protection comparable to that of WldS neurons. Thus, extranuclear WldS expression, most likely due to protein interaction between WldS and the cytoplasmic VCP protein (Laser et al., 2006), results in sufficient ectopic Nmnat activity to confer axon protection. Although one cannot rule out the additional contribution of gene changes caused by WldS expression (Gillingwater et al., 2006b) toward the protective phenotype, current data suggest that the likely basis of WldS-mediated axon protection is through mistargeting of the normally nuclear Nmnat1 protein and its enzymatic activity to extranuclear, possibly axonal compartments.

How does the WldS protein mediate axon protection, and where does this activity intersect with the normal degenerative process? Interestingly, both WldS and Nmnat2 share the same enzymatic domain for NAD+ synthesis (Sorci et al., 2007) that is required for axon protection (Jia et al., 2007; Yan et al., 2010). And as endogenous Nmnat2 activity is essential for axonal survival, it is suggested that the WldS protein protects axons by augmenting or substituting for Nmnat2 to maintain sufficient levels of Nmnat enzyme activity in the distal axons after injury (Gilley and Coleman, 2010). Consistent with this hypothesis, in vivo tracing of GFP-tagged WldS protein in uninjured nerves shows that WldS is normally present in axonal regions where Nmnat2 is also expressed. Moreover, Nmnat2 is found to rapidly degrade, even in WldS nerves, within 4 h after nerve injury, whereas the WldS protein remains stable in the distal axon (Gilley and Coleman, 2010). These reports strongly argue that a molecular mechanism by which the WldS protein confers axon protection is by sustaining key levels of Nmnat activity in the axon that would normally diminish from decreasing Nmnat2 expression after injury (Fig. 3).

Figure 3.
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Figure 3.
A molecular model of WldS-mediated axon protection. The WldS fusion protein, consisting of the full-length Nmnat1 and the first 70 amino acids of Ube4b, is predominantly localized in the nucleus; however, it is also expressed in axonal cytoplasm and organelles such as mitochondria (broken arrows denote known neuronal sites of WldS expression) likely due to interaction with the cytoplasmic VCP protein. Expression of either WldS or extranuclear forms of Nmnat1 is sufficient to protect axons from degeneration upon injury, and this may result from substituting for the activity of Nmnat2 protein, which is degraded quickly after nerve injury. The WldS protein may also augment the enzymatic activity of Nmnat3, a mitochondrial Nmnat isoform, to confer axon protection. The combined result of ectopic Nmnat activities in WldS neurons may be less intracellular Ca2+ release from axoplasmic reticulum or greater Ca2+ buffering by the mitochondria via increased NAD+ production in these organelles, leading to overall decrease in intra-axonal Ca2+ levels (pink arrows denote net direction of Ca2+ flux).

Moreover, as additional Nmnat isoforms exist in different subcellular regions, WldS protein may also promote axonal survival by augmenting the activity of other Nmnat enzymes. Consistent with this, overexpression of Nmnat3, a mitochondrial Nmnat isoform (Berger et al., 2005), results in robust axon protection from traumatic injuries comparable to that of WldS expression in both mammalian and Drosophila models (Avery et al., 2009; Sasaki et al., 2009; Yahata et al., 2009). Although Nmnat3 is not required for normal maintenance of axonal survival (Gilley and Coleman, 2010), its overexpression results in significantly stronger axonal protection than from overexpression of Nmnat2, though this may be due to the labile nature of Nmnat2 protein (Gilley and Coleman, 2010). As the WldS protein is also identified in the mitochondria (Yahata et al., 2009), it raises the notion that WldS expression increases Nmnat activity at multiple intracellular sites to delay axonal degeneration. Assessing whether expression of WldS continues to protect axons in Nmnat2 or Nmnat3 knockouts will help reveal the critical site of Nmnat activity for conferring axonal protection.

Yet, precisely how increased Nmnat enzymatic activity in WldS, Nmnat2, or Nmnat3 protects the axons remains a mystery. As all three proteins contain the highly conserved catalytic domain for the synthesis of NAD+ (Berger et al., 2005), the common metabolite of Nmnat enzyme activities, NAD+ has emerged as an attractive molecular mediator of WldS axon protection. Indeed, exogenous NAD+ is sufficient to protect axons in vitro (Wang et al., 2005; Sasaki et al., 2009). However, this axon protection is only observed at extracellular concentrations above 1 mM, which far exceeds physiological levels, although this may be due to saturation or limited uptake of extracellular NAD+ by putative NAD+ channels on the surface membrane (Alano et al., 2010). Surprisingly, no appreciable difference in NAD+ levels is detected between WldS and wild-type neurons (Mack et al., 2001), and globally reducing neuronal NAD+ levels by chemically inhibiting NAMPT, an enzyme that synthesizes the precursor to NAD+, does not abate WldS axon protection in vitro. Although inhibiting NAMPT activity partially abolishes WldS protection in vivo (Conforti et al., 2009), whether this is due to toxicity from prolonged drug exposure cannot be ruled out. Conversely, increasing overall NAD+ levels in the neuron by blocking PARP and CD38, both NAD+ consuming enzymes, also fails to protect wild-type axons after injury (Sasaki et al., 2009).

A possible explanation for this paradoxical lack of axon protection when cytoplasmic NAD+ levels are raised is that axonal survival requires high NAD+ levels within specific regions or organelles in the axon. Indeed, both Nmnat2 and Nmnat3 are primarily restricted to neuronal ER/Golgi and mitochondrial compartments, respectively (Berger et al., 2005), suggesting that these intra-axonal compartments and membranous organelles may regulate NAD+ concentrations independently from the cytoplasmic NAD+ pool. The presumed effect of a local increase in NAD+ at these intracellular sites in the axon is unclear, though ATP levels drop precipitously after nerve injury, and exogenous NAD addition sustains neuronal ATP levels (Wang et al., 2005). This suggests that rising NAD+ levels may exert axonal protection through increased local energy production. However, Press and Milbrandt (2008) recently reported that increasing mitochondrial Nmnat3 activity is sufficient to delay axon degeneration even from rotenone, a blocker of mitochondrial oxidative phosphorylation. This axonal protection is independent of ATP levels as the rate of ATP loss is similar between Nmnat3-expressing and wild-type axons treated with rotenone (Press and Milbrandt, 2008). Moreover, neither lowering neuronal ATP levels by adding deoxyglucose, an inhibitor of glycolytic enzymes, nor raising ATP levels through TTX, which attenuates ATP consumption by Na/K ATPase, affects axon survival, indicating that maintenance of ATP is unlikely to be the principal mechanism by which increased NAD+ levels promote axon protection. Alternatively, as both the ER/Golgi and mitochondria are capable of sequestering Ca2+, an attractive hypothesis is that sufficient NAD+ levels in these organelles may directly regulate Ca2+ balance in the axon. Indeed, aside from serving as cofactor in oxidative phosphorylation, NAD+ is also capable of modulating ion channel opening (Tamsett et al., 2009). This indicates the possibility that local increase in Nmnat activity, and therefore increase in NAD+ levels, may lower intra-axonal Ca2+ concentrations and protect axons by increasing Ca2+ buffering capacity of mitochondria or decreasing intracellular release by axoplasmic Ca2+ storage sites (Fig. 3). Further work in analyzing differences in Ca2+ flow in subcellular axonal compartments between WldS and wild-type neurons, as well as identifying NAD+-interacting proteins that may gate Ca2+ movement in the axon will be critical in identifying the relationship between Ca2+ regulation and Nmnat activity in promoting axonal survival.

Concluding remarks
Previous studies have independently established the requirements for axonal Ca2+ rise, activities by the ubiquitin–proteasome system and intracellular proteases in axonal degradation. However, how each molecular component interacts to orchestrate the initiation and execution of axonal self-destruction after injury is unclear. A critical insight is gleaned from an early study by Lubińska et al. (1982), who observed that the latency period before the distal axons degenerate is lengthened when the axonal transection is made closer to the cell body. Lubińska and colleagues reasoned that a survival factor is maintained at a critical level in the axon, and when an injury occurs close to the cell body there is a greater level of the trophic factor remaining in the larger distal stump than when the injury occurs more distally. Building upon Lubińska’s initial hypothesis, an appealing unifying model of axon degeneration is that after axonal injury, there is impaired axoplasmic delivery of an axonal survival factor from the soma, which along with continual turnover of the survival factor by the proteasome causes its expression to fall below a critical threshold in the axon. This decreased survival factor activity is sensed by the axon, perhaps via decreased local levels of NAD+, to trigger an execution signal or program that increases intra-axonal Ca2+ levels due to extracellular influx or intracellular Ca2+ release, which then initiates Ca2+-dependent cytoskeletal breakdown (Fig. 2). Such a trophic delivery model helps explain many features of Wallerian axon degeneration, including the physiological latency due to the turnover rate of the survival factor, and the injury-specific directionality of axon degeneration due to the predicted time course of survival factor loss after disrupted transport. It also helps explain the mechanism of WldS protection as one of sustaining Nmnat activity, and thus NAD+ levels above critical threshold in axonal compartments (Fig. 3). The recent identification of the labile molecule Nmnat2, whose endogenous levels in the axon are influenced by proteasomal turnover and whose enzymatic activity is necessary for axon survival, provides exciting experimental support for the survival factor hypothesis.

This survival factor model is also useful for guiding future experiments to answer several key unanswered questions. First, is NAD+, the common enzymatic product of Nmnat2, Nmnat3, and WldS proteins, the critical molecule that mediates axon survival? Identifying the putative molecular targets of NAD+ in the axon, as well as determining whether other metabolites are produced by Nmnat through a metabolomics screen, provide complimentary approaches to identify the key molecular events downstream of Nmnat activity responsible for maintaining axon survival. Second, what are the intermediate events between Nmnat activity and physical proteolysis of the axon? Although direct gating of intra-axonal Ca2+ levels by Nmnat or even NAD+ is appealing in its simplicity, it has yet to be demonstrated experimentally. Finally, do other pathways known to modulate axon degeneration, including several of the “death signals,” interact or converge with Nmnat activity? Interestingly, increased Nmnat activity protects against both Wallerian degeneration and axon degradation from trophic withdrawal (Vohra et al., 2010), suggesting that the final commitment to irreversible axon degeneration may intersect at the levels of Nmnat activity in the axon. Thus, addressing the downstream effectors of WldS/Nmnat-mediated axon protection will be instrumental in identifying therapeutic targets to protect the white matter from many insults besides traumatic injuries. With a working molecular model, one may begin to answer these questions and accelerate the process of identifying the full molecular pathways that underlie axon degeneration in injury and disease.

Back to TopAcknowledgments

We thank Mariko Howe, Mauricio Vargas, and Tom Clandinin for critical reading of the manuscript. The authors apologize to colleagues whose work was not included due to space restrictions.

This work was supported by National Institutes of Health grant EY11310 and the Adelson Medical Research Foundation. J.T. Wang was supported by a Howard Hughes Medical Scientist Research Training Grant and an American Heart Association Pre-doctoral Fellowship.

Back to TopFootnotes

Abbreviations used in this paper:
acute axonal degeneration
Wallerian degeneration slow
Submitted: 18 August 2011
Accepted: 12 December 2011
This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at

Back to TopReferences

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Figure 1.
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Figure 1.
Course of Wallerian axon degeneration. As early as 5–30 min after nerve injury, the axonal segments proximal (left) and distal (right) to the injury site exhibit short-distance acute axon degeneration (AAD), an event that is principally mediated by extracellular Ca2+ influx and activation of the intracellular Ca2+-dependent protease calpain. This event is followed by a slower axonal retraction and formation of axonal bulbs at the injury sites (arrowheads). For the next 24 to 48 h after injury there is a period of relative latency in which the distal axon remains morphologically stable and electrically excitable. Although beading occurs along the distal axon at irregular intervals, there are few signs of physical fragmentation. At more than 72 h after injury, rapid fragmentation and cytoskeletal breakdown occur along the full length of the distal axon, followed by increased glial (consisting primarily of astrocytes, macrophages and, in the PNS, Schwann cells) influx to clear axonal remnants (blue circles) and to possibly promote regenerative attempts by the proximal axon.

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Figure 1.
View larger version:
In this page In a new window
Download as PowerPoint Slide
Figure 1.
Course of Wallerian axon degeneration. As early as 5–30 min after nerve injury, the axonal segments proximal (left) and distal (right) to the injury site exhibit short-distance acute axon degeneration (AAD), an event that is principally mediated by extracellular Ca2+ influx and activation of the intracellular Ca2+-dependent protease calpain. This event is followed by a slower axonal retraction and formation of axonal bulbs at the injury sites (arrowheads). For the next 24 to 48 h after injury there is a period of relative latency in which the distal axon remains morphologically stable and electrically excitable. Although beading occurs along the distal axon at irregular intervals, there are few signs of physical fragmentation. At more than 72 h after injury, rapid fragmentation and cytoskeletal breakdown occur along the full length of the distal axon, followed by increased glial (consisting primarily of astrocytes, macrophages and, in the PNS, Schwann cells) influx to clear axonal remnants (blue circles) and to possibly promote regenerative attempts by the proximal axon.

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‘Teleportation’ is a reality now
PTI | May 4, 2015, 04.07AM IST

LONDON: Ever felt like watching yourself from the sidelines? Scientists have just replicated that feeling in the lab. Neuroscientists have created an out-of-body illusion in people placed inside a br8ain scanner and used that illusion to perceptually ‘teleport’ the participants to different locations in a room.

Scientists from Sweden’s Karolinska Institutet have shown that the perceived location of the bodily self can be decoded from activity patterns in specific brain regions. Studies in rats have shown that specific regions of the brain contain GPS-like ‘place cells’ that signal the rat’s position in the room – a discovery that was awarded the 2014 Nobel Prize in physiology or medicine.
To date, however, it remains unknown how the human brain shapes our perceptual experience of being a body somewhere in space, and whether the regions that have been identified in rats are involved in this process.

In a study published in the scientific journal Current Biology, the scientists created an out-of-body illusion in fifteen healthy participants placed inside a brain scanner. In the experiment, the participants wore head-mounted displays and viewed themselves and the brain scanner from another part of the room.

From the new visual perspective, the participant observes the body of a stranger in the foregr8ou8nd while their physical body is visible in the background, protruding from the bore of the brain scanner.

To elicit the illusion, the scientist touches the participant’s body with an object in synchrony with identical touches being delivered to the stranger’s body, in full view of the participant. “In a matter of seconds, the brain merges the sensation of touch and visual input from the new perspective, resulting in the illusion of owning the stranger’s body and being located in that body’s position in the room, outside the participant’s physical body,” said Arvid Guterstam, lead author of the study.

In the most important part of the study, the scientists used the out-of-body illusion to perceptually ‘teleport’ the participants between different places in the scanner room. They then employed pattern recognition techniques to analyse the brain activity and show that the perceived self-location can be decoded from activity patterns in specific areas in the temporal and parietal lobes.

The scientists could demonstrate a systematic relationship between the information content in these patterns and the participants’ perceived vividness of the illusion of being located in a specific out-of-body position.

“The sense of being a body located somewhere in space is essential for our interactions with the outside world and constitutes a fundamental aspect of human self-consciousness,” Guterstam said.

About garyskeete

ASHWORTH MEDICINE-Professional Medical Assisting, Doctor of Science,Legal Assistant Diploma BSc Criminal Justice PhD Computational Neuroscience MD DSC Epigenetics
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