Background
Injury to peripheral nerves produces abrupt tissue damage at the lesion site where physical impact occurs. The type of trauma determines the extent of damage. For example, compression leads to nerve crush that severs axons but largely preserves the continuity of the nerve connective tissue structures whereas cut and avulsion form a gap between proximal and distal nerve stumps. The nerve segment located distal to and beyond the lesion site undergoes Wallerian degeneration though it does not encounter the physical trauma directly [
1]. Among features that characterize Wallerian degeneration and directly relate to our study, axons disintegrate, myelin sheaths that surround axons break down, a more sustained presence of numerous macrophages replaces an initial short-lived presence of fewer neutrophils, and mostly macrophages and Schwann cells scavenge myelin debris, reviewed in [
2,
3].
To regain function, severed axons must reach the denervated target cells by crossing the lesion site and then entering and growing throughout the Wallerian degenerating nerve segment. Though possible, less than 50% of patients regain adequate sensory and motor functions [
4,
5]. Thus, successful recovery of functions from nerve injury remains an unresolved clinical issue.
Two major factors that affect functional recovery are the type of trauma and the nature of Wallerian degeneration. The type of the trauma determines if and how many of the severed axons successfully cross the lesion site. For example, nerve crush favors successful crossing since the nerve’s uninterrupted connective tissue structures serve as conduits. In contrast, the gap between proximal and distal nerve stumps formed by nerve cut and avulsion injuries obstruct crossing, discussed in detail regarding human nerve injury [
4,
5].
Wallerian degeneration follows all types of nerve injury. Our current study relates to the question how does Wallerian degeneration affect the growth of those severed axons that successfully crossed the lesion site and thereby the recovery of function. In this context, at least three factors together determine whether severed axons successfully grow/regenerate throughout the Wallerian degenerating nerve segment. First is the length of the distal nerve segment. It can vary from several millimeters to up to over 1 m depending on species (e.g., mice versus humans) and site of trauma (e.g., near to versus distant from denervated target cells). Second is the slow growth of axons, which prolongs the time it takes axons to regenerate long distances. Third is the decline with time of the capacity to support axon growth that initially develops in Wallerian degeneration. Observations in humans and studies in experimental animals support this understanding. In humans, the rate that severed axons regenerated through Wallerian degenerating nerve segments was examined at different times after nerve crush injuries that did not require surgical reconstruction and after more extensive nerve injuries that required suturing of the proximal and distal nerve stumps. Human sensory axons that regenerated at an initial rate of 2.5 mm/day slowed down to 0.5 mm/day over a period of 200 days and human motor axons that initially regenerated 2 mm/day slowed down to 1 mm/day [
6,
7]. In nonhuman primates, fast regeneration was the major determinant of successful recovery [
8]. In rats, axon growth support, which initially developed in Wallerian degeneration, diminished with time, resulting in poor recovery [
9,
10].
A leading cause of slow axon regeneration in the Wallerian degenerating nerve segment is myelin debris (also referred to as degenerated myelin in the literature). Myelin debris and MAG (myelin-associated glycoprotein) inhibited axon growth in culture and in vivo [
11‐
14]. Schwann cells and macrophages could alleviate axon growth inhibition since the two scavenge myelin debris in Wallerian degeneration. Schwann cells removed myelin debris by autophagy [
15,
16] and phagocytosis [
16,
17], and macrophages by phagocytosis [
18‐
20]. However, the rate of myelin debris removal is very likely slower than the rate fast regenerating axons grow. This is suggested by live imaging observations that revealed the presence of residual myelin debris (i.e., myelin debris that was not yet removed) confronting and slowing down the growth of regenerating axons [
21].
Myelin debris itself and serum can impede the removal of myelin debris in Wallerian degeneration. Previously, we showed that myelin debris inhibited its own phagocytosis in cultured primary macrophages and microglia through the binding of CD47 on myelin to the immune inhibitory receptor SIRPα (also known as CD172a, SHPS-1, p84, gp93, and BIT) on macrophages and microglia [
22,
23]. In turn, SIRPα generated “don’t eat me” signaling, which reduced phagocytosis. We further showed that serum triggered SIRPα-dependent inhibition of myelin debris phagocytosis. As serum components are present in tissues other than central nervous system [
24], serum may also exist in peripheral nerves. Therefore, not only CD47 on myelin but also serum may trigger SIRPα-dependent inhibition of myelin debris removal.
Yet, it remained unclear whether SIRPα-dependent inhibition of myelin debris phagocytosis impedes the removal of myelin debris in vivo, leading residual myelin debris to slow down axon regeneration and so delay the recovery of functions. To resolve this issue, our present study used SIRPα null (SIRPα−/−) mice and littermate wild-type (SIRPα+/+) mice [
25] to analyze myelin debris removal, axon regeneration, macrophage presence and phagocytic capacity, and the recovery of sensory and motor functions. Our findings suggest that SIRPα-dependent inhibition of myelin debris phagocytosis in macrophages impedes the removal of myelin debris in Wallerian degeneration, leading to slow axon regeneration and so to delayed recovery of functions from peripheral nerve injury.
Methods
Animals
C57BL/6 J mice from which SIRPα was deleted (SIRPα null; homozygous SIRPα−/−) and heterozygous SIRPα+/− were generated at Kobe University Graduate School of Medicine, Japan [
25]. Heterozygous SIRPα+/− were used to establish SIRPα−/− and littermate wild-type/SIRPα+/+ C57BL/6 J mice colonies at the Hebrew University Faculty of Medicine animal facility. Sex- and age-matched 8- to 12-week-old mice were used in experiments in accordance with the Israeli national research council guide for the care and use of laboratory animals and the approval of the Hebrew University institutional ethic committee.
Surgical procedures
Surgery was performed under anesthesia on one hind limb of wild-type and SIRPα−/− mice. Sciatic and saphenous nerves were exposed through small incisions in the overlaying skin. Freeze-crush injuries that enable axon regeneration were performed on sciatic and saphenous nerves using a fine jeweler’s tweezer that was cooled in liquid nitrogen and then applied to nerves for 5 s, taking care to preserve the continuity of the epineurium. Avulsion injuries that do not enable axon regeneration were performed on sciatic nerves by removing a small nerve segment at mid-thigh level. Finally, the skin was sutured and sprayed with antiseptics.
Assessment of functional recovery after nerve injury
Two investigators assessed the recovery of sensory and motor functions independently by testing all wild-type and SIRPα−/− mice side by side at 1-day intervals after surgery. Each mouse was tested for at least 3 days after function first returned to verify consistency.
Recovery of sensory function was assessed using the flexion-withdrawal reflex: withdrawal of hind limbs in response to touching their paws with a blunt pin and von-Frey monofilaments that produce punctate mechanical stimuli delivered mostly by Aδ axons, i.e., pinprick testing [
26]. Mice that had their saphenous nerve freeze-crushed were placed on an elevated wire mesh platform until calm, and then, testing of both injured and uninjured limbs was carried out by gently touching paws at areas that saphenous sensory axons normally innervate.
Recovery of motor function was assessed using the toe-spreading reflex: spreading of the toes in response to gently lifting mice by their tail. The reflex was tested in both the injured and uninjured hind limbs.
Preparation of BMDM (bone marrow-derived macrophage)
We followed previously published protocols [
27‐
29] with some modifications. Femur and tibia bones were removed from wild-type and SIRP−/− mice and placed in complete MEM supplemented with 15% heated inactivated FCS, 2 mM glutamine, MEM non-essential amino acids, MEM vitamin solutions, 1 mM sodium pyruvate, 1ug/ml transferrin APO, 100 U/ml penicillin, and 100 mg/ml streptomycin (Biological Industries, Beit Haemek, Israel). Bone marrow was flushed out, cells suspended in red blood cell lysis buffer for 1 min, washed in complete MEM, and plated in cell culture petri dishes for 2 to 4 h at 37 °C. Non-adherent bone marrow-derived cells that include macrophage precursor cells were plated in 100 mm plastic/bacteriological dishes (0.4 10
6 cells/dish) in complete MEM supplemented with 15% L929-cell conditioned medium that contains the macrophage MCSF (colony-stimulating factors) [
30]. Macrophage precursor cells that differentiated into adherent BMDM after one week in the presence L929 cells conditioned media were used in experiments.
Myelin isolation
The detailed protocol for isolating myelin was previously described [
31]. Isolated myelin is “myelin debris” since intact myelin breaks down during isolation.
Phagocytosis of myelin debris
Phagocytosis was assayed as previously described [
31]. BMDM were plated in 96-well tissue culture plates at a density that minimizes cell-cell contact in the presence of DMEM supplemented by 10% FCS. Non-adherent BMDM were washed out after 2 h and adherent BMDM left to rest overnight. Next, BMDM were washed and myelin debris added in DMEM/F12 in the presence of serum for 40 min, unphagocytosed myelin debris washed out, and levels of phagocytosis determined by ELISA. At this time all myelin debris was phagocytosed/internalized [
31,
32].
Detecting and quantifying myelin debris phagocytosis by ELISA
This assay is based on the detection of the myelin-specific protein MBP (myelin basic protein) in phagocyte as previously detailed [
31]. Since MBP is unique to myelin and BMDM do not produce it, MBP levels in BMDM cytoplasm are proportional to levels of phagocytosed myelin debris. In brief, BMDM were immediately lysed (50 mM carbonate buffer, pH 10) after myelin debris phagocytosis was completed, and lysates transferred to high protein absorbance plates (Thermo Fisher Scientific, Nunc International, USA) in equal volume of coating buffer (0.5 M carbonate buffer pH 9.6). Levels of MBP were determined by ELISA using rat anti-MBP mAb and matching control IgG (Bio-Rad Laboratories Inc., Hercules, USA).
When phagocytosis by BMDM from wild-type mice was compared with phagocytosis by BMDM from SIRPα−/− mice, phagocytosis by each population was first normalized to the number of respective BMDM counted in 1-mm2 areas at the center of wells. Normalizing phagocytosis to cell number is required since BMDM from the two strains of mice may differ in their adherence properties, thus resulting in different number of adherent cells even when the same number of cells was initially seeded. To this end, BMDM in replicate plates were fixed, stained, and counted. Phagocytosis by BMDM from SIRPα−/− mice was calculated as percentage of phagocytosis by BMDM from wild-type mice normalized to 100%.
Quantifying MBP content in nerve tissue
The detailed protocol used to quantify Galectin-3/MAC-2 [
27] was previously adopted to quantify MBP in peripheral nerves [
33]. In brief, nerves were homogenized in 50 mM sodium carbonate buffer pH 9.0 supplemented with protease inhibitor cocktail (Sigma-Aldrich, Saint Louis, USA), and protein concentration in cleared extracts was determined using the Bradford assay reagent (Bio-Rad Laboratories Inc., Hercules, USA) and adjusted to 5 μg/mL. Equal volumes (75 μL) of extracts and coating buffer (0.5 M carbonate buffer pH 9.6) were incubated overnight at 4 °C in 96-well high protein absorbance plates (Thermo Fisher Scientific, Nunc International, USA), and levels of MBP determined by ELISA using rat anti-mouse MBP mAb (Bio-Rad Laboratories Inc., Hercules, USA).
Immunoblot analysis
BMDM were plated in 10 cm tissue culture plates at a non-confluent density (3 × 106 cells per plate) in the presence of DMEM supplemented by 10% FCS and left to rest overnight. BMDM were washed in PBS and lysed in ice cold lysis buffer (50 mM Tris HCL pH 7.5, 20 nM MgCl2, 150 mM NaCl, 0.5% NP-40, 10 μM DTT, and 100 μM NaVa) supplemented with protease and phosphatase inhibitors cocktail (Sigma-Aldrich, St-Louis, USA) and total protein content determined using Bradford reagent. Equal protein content from whole cell lysates was separated on SDS-PAGE. Then, proteins were blotted to nitrocellulose membranes, blocked in 5% BSA in TBST (Tris-buffered saline supplemented by 0.1% Tween 20) for 1 h at RT, and incubated over night at 4 °C in the presence of primary antibodies rabbit anti-SIRPα (Abcam, Cambridge, UK) and mouse anti-α-tubulin (Sigma-Aldrich, St-Louis, USA). Blots were washed with TBST and incubated with respective secondary antibodies goat anti-rabbit and goat anti-mouse conjugated to HRP (Jackson Immuno Research, USA) for 40 min at RT. Proteins were visualized by enhanced chemiluminescence for HRP detection.
Immunofluorescence microscopy
Nerves were cross-sectioned (15 μm) in a freezing cryostat, sections fixed in 4% paraformaldehyde in PBS for 15 min, washed 3x10 min in PBS containing 0.1% triton (PBST), blocked (10% normal donkey serum, NDS, in PBST) for 1 h at room temperature, and incubated overnight at 4 °C in the presence of primary antibodies as described below. To visualize CD47 and SIRPα, sections were incubated in rat anti-mouse CD47 and SIRPα monoclonal antibody (mAb) (provided by Dr. Oldenborg, Umea, Sweden). To visualize macrophages, sections were incubated in hybridoma supernatants containing rat anti-mouse mAb M1/70 (Developmental Studies Hybridoma Bank, Iowa City, USA) and mAb 5C6 (American Type Culture Collection, Rockville, USA) against αM/CD11b subunit of CR3 (complement receptor-3). To visualize SIRPα in macrophages, sections were double stained with mAbs against αM/CD11b subunit of CR3 (see above) and anti-SIRPα Ab8120 (Abcam, Cambridge, UK). To visualize axons, sections were incubated in anti-neurofilament (Sigma-Aldrich, Saint Louis, USA) (diluted 1:200 in 2% NDS in PBST). The following day, sections were washed 3x10 min in PBS and incubated with the corresponding secondary antibodies: donkey anti- rabbit CY3 1:1000 and FITC-conjugated rabbit anti-rat IgG (Jackson IR laboratories, PA, USA) in 2% NDS for 1 h at room temperature, washed 3X10min in PBS, stained with Hoechst (1:2000) for 2 min, and washed in PBS for 10 min. Finally, sections were air-dried, mounted on coverslips, and sealed with Fluoromount (Sigma-Aldrich, Saint Louis, USA). All steps involving fluorescence were carried out in a slightly darkened room. The immuno-stained slices were examined under a fluorescence microscope.
Statistical analysis
The following statistical analyses were carried out using GraphPad Prism software: Gaussian distribution, the parametric unpaired t test and one- and two-way ANOVA, the nonparametric Mann-Whitney test, and the log-rank Mantel-Cox test. Data that passed the normality test were subjected to parametric statistics and those that were too small for testing for normality were subjected to nonparametric statistics.
Discussion
This study is the first to reveal an intrinsic normally occurring SIRPα-dependent mechanism that impedes the in vivo removal of myelin debris by inhibiting phagocytosis in macrophages present in Wallerian degeneration, leading to slow axon growth/regeneration through Wallerian degeneration and thereby to delayed recovery of function from nerve injury. We suggest this mechanism based on the following major results and arguments. First, compared with wild-type/SIRPα+/+ mice, deletion of SIRPα increased the phagocytic capacity of macrophages and further accelerated the removal of myelin debris in SIRPα−/− mice. Since macrophages and Schwann cells scavenge most of myelin debris in Wallerian degeneration, and only macrophages express SIRPα in wild-type/SIRPα+/+ mice, deletion of SIRPα could only have taken place in macrophages and not in Schwann cells. Therefore, the accelerated removal of myelin debris in SIRPα−/− mice is most likely due to the increased phagocytic capacity of SIRPα−/− macrophages. Second, compared with wild-type/SIRPα+/+ mice, deletion of SIRPα accelerated axon regeneration through Wallerian degenerating nerve segments and so facilitated the recovery of functions in SIRPα−/− mice. The faster removal of axon growth-inhibitory myelin debris in SIRPα−/− mice accounts most likely for faster axon growth. This is suggested by the findings in wild-type/SIRPα+/+ mice that the slower removal of myelin debris was associated with slower axon growth and delayed recovery, supported by live imaging of myelin debris slowing axon growth [
21]. All those findings together suggest that in SIRPα−/− mice, deletion of SIRPα increases the phagocytic capacity of macrophages, leading to faster removal of axon growth-inhibitory myelin debris, and so to accelerated axon growth and facilitated recovery of functions. By contrast, in wild-type/SIRPα+/+ mice, SIRPα impedes the phagocytic capacity of macrophages, leading to protracted removal of myelin debris that slows axon growth, and so prevents severed axons from fully implementing their regenerative potential.
The role that we suggest to assign to SIRPα in the recovery of function from nerve injury requires that genetic deletion of SIRPα increases the phagocytic capacity of SIRPα−/− macrophages by eliminating SIRPα-dependent inhibition of phagocytosis and not by inducing another mechanism. Our previous findings in cultured primary wild-type/SIRPα+/+ macrophages and microglia suggest that this is the case. Function blocking antibodies against SIRPα and its ligand CD47, phagocytosis of myelin debris that does not express CD47, and knocking down SIRPα all augmented phagocytosis [
22,
23]. Since both genetic deletion of SIRPα and the genetic-independent prevention of SIRPα activation (i.e., blocking SIRPα-CD47 binding and eliminating CD47 from myelin debris) augmented phagocytosis, the increased phagocytic capacity is most likely due to the elimination of SIRPα-dependent inhibition of phagocytosis and not by another mechanism that genetic deletion of SIRPα may have induced.
Another point of consideration is whether deletion of SIRPα facilitated axon growth and the recovery of function by preventing SIRPα from directly inhibiting axon growth by acting on or in neurons/axons and not indirectly by preventing SIRPα from impeding the removal of axon growth-inhibitory myelin debris, which we suggest. Experimental findings in mice that express SIRPα support our proposition and not the alternative. For example, the deletion of axon growth-inhibitory MAG from myelin in mice that also display exceptionally slow removal of myelin debris advanced the growth of severed axons in the presence of myelin debris [
13], and further, axons grew faster through Wallerian degenerating nerve segments from which myelin debris had previously been removed (discussed below under “conditioning injury”).
Neutrophils, which infiltrate the Wallerian degenerating nerve early but transiently after injury, could phagocytose and so contribute to the removal of myelin debris [
37,
38]. Two scenarios warrant consideration in this regard. First, neutrophils, as macrophages, express both the phagocytic receptor CR3 and inhibitory SIRPα [
39]. Thus, deletion of SIRPα should augment phagocytosis in neutrophils as in macrophages. Second, it is most probable that the overall contribution of macrophages to the phagocytosis of myelin debris is considerably greater than that of neutrophils since the presence of neutrophils in Wallerian degeneration is transient, and further, macrophages outnumber neutrophils exceedingly through the entire period of myelin debris removal. We reached this understanding based on the findings that neutrophils were outnumbered by macrophages 3 to 1 on day 3 after injury, and then, at day 7 after injury, neutrophils were practically absent [
38]. In contrast and during the same period, the number of macrophages (i.e., cells expressing the murine macrophage specific F4/80 antigen) increased progressively [
17,
35‐
37]. Thus, in Wallerian degeneration, a sustained presence of numerous macrophages replaces an initial short-lived presence of substantially fewer neutrophils through the entire period of myelin debris removal.
Our proposition that macrophages phagocytosed exceedingly more myelin debris than neutrophils seemingly contradicts the finding that deletion of neutrophils reduced myelin debris removal [
38]. This apparent discrepancy can be reconciled if neutrophils contributed to macrophage recruitment in Wallerian degeneration as they do in acute inflammation in other tissues. In acutely inflamed tissues, an initial transient infiltration of short-lived neutrophils was replaced by a sustained presence of macrophages [
40,
41]. This transition was governed by the constitutive apoptotic death of the infiltrating neutrophils and the associated shedding of IL6R (interleukin-6 receptor) from their surface, leading to IL6 trans-signaling that inhibited the production of leukocyte chemoattractants and at the same time induced the production of monocyte chemoattractants. In agreement with that, depletion of neutrophils resulted in reduced levels of soluble IL6R and reduced accumulation of macrophages in inflamed tissues, confirming that normally the initial infiltration of neutrophils prompted macrophage recruitment [
41]. We previously showed that fibroblasts produce significant levels of IL6 in Wallerian degeneration within hours after nerve injury, and further, macrophages also produce IL6 but at later stages [
42]. The early production of IL6 in Wallerian degeneration [
42] and the early recruitment of neutrophils to Wallerian degeneration [
38] together suggest that neutrophils could have contributed to macrophage recruitment in Wallerian degeneration as they do in acute inflammation [
40,
41].
Our findings highlight the fact that fast removal of myelin debris is critical to recovery from nerve injury and, further, that the contribution of macrophages to the process is significant. In agreement with this understanding are the observations that excluding macrophages from Wallerian degeneration reduced the removal of myelin debris and impaired recovery [
43‐
45] and, further, that the slower removal of myelin debris delayed axon regeneration and muscle reinnervation in aged mice compared with young mice. The removal of myelin debris adds to additional innate-immune functions significant to recovery that macrophages carry out. For example, we previously documented that macrophages in Wallerian degeneration were poor producers of the inflammatory cytokines TNFα (tumor necrosis factor-α), IL1β and GM-CSF (granulocyte-macrophage colony-stimulating factor) [
27,
46]. In contrast, the same macrophages were major producers the cytokines IL10 and IL6 [
33,
42]; IL10 is a major anti-inflammatory cytokine [
47] and IL6 has context-dependent anti-inflammatory and inflammatory properties [
48,
49] as well as neurotrophic properties [
50]. Thus, macrophages in Wallerian degeneration are mostly of the M2 tissue repair phenotype, reviewed and discussed in [
2,
3,
51‐
53]. Interestingly, a transient short-lived infiltration of neutrophils preceded the infiltration of macrophages that then acquired the M2 phenotype at sites of traumatic injury to spinal cords [
54].
The rate of myelin debris removal may affect fastest growing axons most and slower growing axons less or not at all. We base this suggestion on the time course of myelin debris removal in wild-type mice shown here and previously [
33] and the fact that the rate of axon growth can vary over a wide range; e.g., from 0.28 to 2.16 mm/day [
21]. Hence, fastest growing axons may precede debris removal and debris removal may precede slowest growing axons. Consequently, fastest growing axons may confront more residual myelin debris than slower growing axons and slowest growing axons may not confront myelin debris at all.
It is of interest to interpret previous findings by others in light of our current results and understanding. Rat axons that regenerated through Wallerian degenerating nerve segments grew slower after a single injury than after a test injury that followed a previously inflicted “conditioning injury”, e.g., [
55]. Sensory axons regenerated 4.02 mm/day after a single injury and 6.76 mm/day after a test injury that followed a conditioning injury inflicted 7 days before [
55]. The authors of this study suggested that cell body changes and the environment of the lesioned axons acted in an additive fashion to produce a maximal conditioning lesion phenomenon. Most studies that followed focused on mechanisms within and surrounding the cell bodies of the axotomized neurons to explain faster regeneration after conditioning lesions, reviewed and discussed in [
52]. Our study suggests that the removal of growth-inhibiting myelin debris from the path through which axons regenerate accelerates axon growth and recovery. Taking the normal time course of myelin debris removal in Wallerian degeneration that we showed here and previously [
33], the 7-day time interval between conditioning and test injuries enabled removing most of the myelin debris before severed axons began growing after the test injury. By contrast, very little of myelin debris was removed by the time regeneration commenced after a single injury. Notably, the conditioning lesion increased the rate rat sensory axons regenerated 1.68-fold [
55]. This is comparable to the 1.4-fold increase in the recovery rate of sensory function caused by removing SIRPα-dependent inhibition of myelin debris removal in mice that we report here since accelerated recovery of function is mostly due to sensory axons regenerating faster.
The speed at which severed axons regenerate through Wallerian degenerating nerve segments affects recovery since axon growth support that initially develops in the Wallerian degenerating nerve tissue declines with time. In rat, recovery deteriorated progressively after delaying regeneration by 8 to 24 weeks and GDNF (glial cell line-derived neurotrophic factor) that Schwann cells initially upregulated declined gradually as of 4 weeks after nerve injury [
9,
56]. In mice, levels of c-Jun that Schwann cells upregulated early after nerve injury dropped significantly at 10 weeks after injury [
57].
A progressive slowdown of axon regeneration through Wallerian degenerating nerve segments occurs in humans both after nerve crush injuries that did not require reconstructive surgery and after more extensive nerve injuries that did. The regeneration of sensory axons slowed from 2.5 mm/day to 0.5 mm/day and the regeneration of motor axons from 2 mm/day to 1 mm/day [
6,
7]. The findings in rodents [
9,
56,
57] raise the possibility that the progressive slowdown of axon regeneration in humans is due to, at least in part, to the gradual time-dependent decline of axon growth support that Wallerian degeneration provides. If that is the case and axon growth-support declines considerably at 10 weeks after injury in humans, then axons regenerating 2 mm/day may face injured nerve tissue with reduced ability to support axon regeneration at a distance of about 14 cm from an injury site (i.e., about half or less the distance from elbow to wrist in adults). Thus, regenerating axons are likely to cover longer distances before axon growth support deteriorates by speeding up their rate of growth through the Wallerian degenerating nerve segment.
Our findings that SIRPα inhibits myelin debris phagocytosis in macrophages present in Wallerian degeneration that follows nerve crush injury is most likely relevant to all types of nerve injury since Wallerian degeneration follows all of them. Our study further suggests that blocking SIRPα-dependent inhibition of myelin debris leads to faster regeneration and so to facilitated recovery of function. However, this mechanism can only affect severed axons that successfully cross the lesion site and then enter the distal nerve segment. In humans, nerve crush injuries, as we used in this study, favor crossing and entry to the distal nerve segment whereas more extensive nerve injuries (e.g., cut and avulsion) require surgical reconstruction to achieve this goal. Unfortunately, depending on the nature of trauma, surgical reconstruction has its limitation, discussed in [
4,
5].
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