Extracellular Pgk1 enhances neurite outgrowth of motoneurons through Nogo66/NgR-independent targeting of NogoA

The inhibition of NOM by muscle-specific NogoA might result from inducing a canonical Nogo66/NgR or Nogo66/PirB signaling pathway or from influencing the secretion of myokines. To define this molecular mechanism, we employed mouse Sol8 myoblasts and established a Doxycycline (Dox)-inducible cell line stably harboring the Sol8-vector only (Sol8-vector) and the Sol8 vector with NogoA insert (Sol8-NogoA) (Figure 1—figure supplement 1). After induction, the CM obtained from cultured myoblast cells harboring Sol8-vector was directly used to culture motoneuron cell line NSC34. After culturing, motoneurons exhibited extended neurites, while motoneurons incubated with Sol8-NogoA CM failed to exhibit neurite outgrowth (Figure 1—figure supplement 1). Importantly, NogoA was not detected in the CM cultured by Sol8-NogoA cells after Dox induction for 48 hr (Figure 1—figure supplement 1C). This line of evidence suggested that the component(s) contained in CM from cultured Sol8-NogoA play(s) a role in NOM inhibition, but not through NogoA contained in medium. Moreover, unlike neurons treated with Sol8-vector, Western blot analysis demonstrated that the signal intensity of phosphorylated Cofilin at S3 (p-Cofilin-S3), which is a hallmark of growth cone collapse in neuronal cells, as well as a marker of reduced axonal actin dynamics in ALS patients with depleted and mutated C9ORF72 (Heredia et al., 2006; Sivadasan et al., 2016), was upregulated in neurons treated with Sol8-NogoA CM (Figure 1—figure supplement 1). Collectively, this line of evidence suggested that the component(s) contained in CM from cultured Sol8-NogoA play(s) a role in NOM inhibition.

We further employed 2D electrophoresis, followed by LC MS/MS, to analyze the total proteins content in CM. Among these examined proteins, we found that the level of Pgk1 protein contained in Sol8-NogoA CM was significantly reduced compared to that in Sol8-vector CM (see Figure 1—figure supplement 2). Furthermore, the degree of p-Cofilin expression in NSC34 cells cultured in CM from Pgk1-overexpressed Sol8-NogoA cells was significantly reduced compared with that in NSC34 cells cultured in CM from Sol8-NogoA cells. Therefore, Pgk1 was chosen for further study to confirm its potential role in NOM. Additionally, we used Western blot analysis to detect the protein level of Pgk1 in the sera of transgenic SOD1 G93A mice, in which NogoA was overexpressed in muscle (Figure 1—figure supplement 3), the results of which were consistent with those reported by Bros-Facer et al. (2014). We found that the amount of Pgk1 in the sera of SOD1 G93A mice was significantly reduced compared with that in the sera of WT mice (Figure 1—figure supplement 3). Collectively, based on the above results provided in vitro and in vivo evidence, we chose Pgk1 for further study to confirm its potential role in NOM.

We added Pgk1 directly into Sol8-NogoA CM and found that p-Cofilin-S3 in NSC34 cultured with Sol8-NogoA CM plus Pgk1 was lower than that of cells cultured with Sol8-NogoA CM (Figure 1A). Importantly, supplementary Pgk1 could reduce p-Cofilin-S3 to then restore NOM of NSC34 cells, even though they had been cultured in Sol8-NogoA CM, indicating that stalled NOM of NSC34 cells cultured in Sol8-NogoA CM could be rescued upon the addition of Pgk1.

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To further confirm whether Pgk1 secreted by muscle cells could enhance NOM, we added Pgk1-specific antibody to Sol8-vector CM and then cultured NSC34 cells. In response, NOM was totally inhibited and p-Cofilin-S3 was higher compared to those observed in the control groups, such as Sol8-vector CM and Sol8-vector CM plus rabbit IgG (Figure 1B). Thus, we suggest that Pgk1 is secreted by myoblasts and that it exists in muscle cell CM to play an important role in enhancing NOM.

Next, to determine if Pgk1 alone is sufficient to induce NOM and neuronal differentiation, we switched to low trophic factor differentiation media (DM). Under the same condition, Pgk1 was able to extend NOM and downregulate p-Cofilin-S3 (Figure 1C). We also found that the effect of adding Pgk1 on the downregulation of p-Cofilin-S3 was dose-dependent from 0, 11, 33, 66 and 99 ng/ml (Figure 1—figure supplement 4). Moreover, when NSC34 cells were treated with Pgk1 at 33 ng/ml for 0, 8, 16, 24 and 48 hr, we found that the effect of Pgk1 on the downregulation of p-Cofilin-S3 was dose-dependent from 8 through 24 hr (Figure 1—figure supplement 4). However, we noticed that p-Cofilin-S3 was unexpectedly increased if cells were treated with Pgk1 for 48 hr. The doses of Pgk1 we examined did not cause any negative effect on cell growth and survival. However, based on the above results and to ensure that we did not seriously affect cell physiology, we treated NSC34 cells with the minimal concentration (33 ng/ml of Pgk1 for 24 hr) throughout the entire study. Interestingly, the addition Pgk1 at that concentration not only rescued the number of neurite-bearing cells, but also enhanced neurite length of NSC34 cells cultured in Sol8-NogoA CM and DM (Figure 1D–E).

To further learn if these restored neurites in NSC34 cells could differentiate into mature neurons, we employed antibody combined with fluorescence staining to specifically detect Syn1, which is a marker labeling neuronal synaptic vesicles in cell terminals (Fletcher et al., 1991), and maturation markers, such as MAP2, GAP43 and ChAT (Maier et al., 2013). Based on the distribution by immunofluorescence staining against Syn1 within NSC34 cells cultured in CM, the percentages of NSC34 cells displaying the Syn1 signal in the growth cones terminal were 43 ± 4.78, 13 ± 1.63 and 45 ± 4.55% in Sol8-vector CM, Sol8-NogoA CM and Sol8-NogoA CM plus Pgk1, respectively (Figure 2A–D). The number of neurons with Syn1 signal at the end of growth cones could be restored to that cultured in Sol8-vector CM after Pgk1 was added in Sol8-NogoA CM.

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We next examined the expression levels of functional proteins, such as MAP2, GAP43, Syn1 and ChAT, in NSC34 cells. The expressions of MAP2, GAP43, Syn1 and ChAT were all increased when Pgk1 was added to the Sol8-NogoA CM/NSC34 culture (Figure 2E), suggesting that Pgk1 addition could rescue weak cell differentiation caused by culturing in Sol8-NogoA CM. Taken together, we conclude that exogenous addition of Pgk1 enables NSC34 cells cultured in Sol8-NogoA CM to improve differentiation, maturation, and neurite growth.

Since Pgk1 is a key enzyme for glycolysis, it is necessary to determine whether reduced p-Cofilin-S3 and resultant enhancement of NOM by addition Pgk1 could be attributed to glycolytic effect. Neither the absence nor presence of ePgk1 resulted in any significant difference in glycolytic function of neurons (Figure 3A). Even in the low glycolytic condition induced by glycolysis inhibitor 3-(3-Pyridinyl)−1-(4-pyridinyl)−2-propen-1-one (3PO) (Figure 3B), in which p-Cofilin-S3 was insignificant compared to the untreated group (Figure 3C, lanes 1 vs. 3), ePgk1 still downregulated p-Cofilin-S3 (Figure 3C, lanes 3 vs. 4). Moreover, addition of Pgk1 mutants, Pgk1-T378P and Pgk1-D375N, both of which lost their catalytic activities (Lay et al., 2002; Chiarelli et al., 2012) in DM, could still reduce p-Cofilin-S3 (Figure 3D), suggesting ePgk1 promotes NOM independent of glycolysis.

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To confirm the role of Pgk1 secreted from muscle cells regulates motoneuron development in vivo, we employed the CRISPR/Cas9 system to knock out Pgk1 in zebrafish embryos. We used muscle-specific α-actin promoter to drive overexpression of Cas9 in muscle cells in order to exclusively knock out the pgk1 gene. Since turbo-red fluorescent protein (tRFP) was engineered to fuse with Cas9 and P2A peptide, it served as a reporter to reflect the overexpression of Cas9 (Figure 4A). Compared to control Tg(mnx:GFP) embryos at 30 hpf (Figure 4C), the tRFP signal was observed in the muscle of pZα-Cas9-injected embryos, indicating that Cas9 was overexpressed in certain muscle cells (Figure 4D). Nevertheless, motoneurons were normally developed, which suggests that overexpression of Cas9 in muscle cells had no effect on development. However, when Tg(mnx:GFP) embryos were coinjected with pZα-Cas9 and pgk1 sgRNA, which inhibits the production of Pgk1 in muscle cells (Figure 4—figure supplement 1), defective motoneurons were observed (Figure 4E), suggesting that the reduction of Pgk1 in muscle cells is followed by impairment of NOM. In a parallel experiment, by conditional knockout of phosphoglycerate mutase 2 (pgam2) in muscle cells (Figure 4—figure supplement 1), a downstream enzyme of Pgk1 in the glycolytic pathway, we again confirmed that the glycolysis pathway in muscle did not affect NOM (Figure 4F). Collectively, these results suggested that defects in NOM result from a decrease in Pgk1 secretion in muscle cells. To prove the consistency of this result in vivo, we constructed a plasmid containing Pgk1 fused with P2A peptide and tRFP (Figure 4B). After injecting this construct into zebrafish embryos, RFP exhibited specifically in muscle cells. When these RFP-expressing cells were sorted out and examined by Western blot analysis, the results demonstrated that Pgk1 was overexpressed (Figure 4—figure supplement 1). Meanwhile, motoneuron axons exhibited ectopic growth (Figure 4G). As suggested by quantification of the axonal extension phenotype shown in Figure 4—figure supplement 2, the increase of Pgk1 in muscle cells enhances NOM.

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We next sought to clarify the molecular pathway that allows addition of Pgk1 to promote neuronal differentiation and NOM of NSC34 cells. The binding of Nogo66 domain to Nogo receptor (NgR) induces an acute intracellular signaling response to inhibit NOM through (i) increasing p-Cofilin-S3 (Ohashi et al., 2000; Manetti, 2012) by phosphorylating Limk1 (p-Limk1-T508) in the Rho/ROCK pathway, (ii) increasing phosphorylated EGFR (p-EGFR-Y1173) (Koprivica et al., 2005), or (iii) downregulating AKT phosphorylation (p-AKT-S473) (Wang et al., 2011). Here, we demonstrated there was no significant changes of p-ROCK2-Y256, p-EGFR-Y1173 or p-AKT-S473 expressions among NSC34 cells cultured in Sol8-vector CM, Sol8-NogoA CM and Sol8-NogoA CM plus Pgk1 (Figure 5—figure supplement 1). Surprisingly, Sol8-NogoA CM plus Pgk1 did reduce p-Cofilin-S3 via p-Limk1-S323, but not p-Limk1-T508 (Figure 5A), suggesting that the ePgk1-mediated pathway is independent of the Nogo66/NgR/ROCK2-Y256/Limk1-T508 axis.

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The overexpression of Limk1, Limk1-S323A and Limk1-T508V in NSC34 cells cultured in DM resulted in increasing the basal level of intracellular p-Cofilin-S3 (Figure 5B). When we treated cells overexpressing Limk1, Limk1-S323A, and Limk1-T508V with Pgk1, the cells overexpressing Limk1 still responded to Pgk1, as evidenced by decreased p-Cofilin-S3 (Figure 5C). Overexpression of Limk1-S323A, but not Limk1-T508V, abolished Pgk1-mediated p-Cofilin-S3 downregulation (Figure 5C), again confirming that Pgk1 decreased p-Cofilin-S3 through Limk1-S323, but not Limk1-T508.

We further identified upstream kinases, including P38/MK2 and Rac1/Pak1 (Figure 5D), along with their phosphorylated sites, using Sol8-vector CM and Sol8-NogoA CM in the absence and presence of Pgk1. Pak1 overexpression confirmed this series of signaling transduction pathway, consistent with our results using Sol8-vector CM and Sol8-NogoA CM (Figure 5E). On the other hand, when NSC34 cells were cultured in DM plus a Pak1 inhibitor, FRAX597, we found that the intracellular p-Pak1 was reduced and that the levels of p-P38-T180, p-MK2-T334, p-Limk-S323 and p-Cofilin-S3 were also reduced (Figure 5—figure supplement 2), suggesting that P38, MK2, Limk, and Cofilin are all downstream effectors of Pak1. In sum, ePgk1 triggers a reduction in p-Cofilin-S3, in turn promoting NOM through decreasing a novel p-Pak1-T423/p-P38-T180/p-MK2-T334/p-Limk1-S323 axis via reducing Rac1-GTP activity in neuronal cells.

To confirm whether Pgk1 involved regulating pathway is independent of activation of NgR receptor by NogoA. First, we determined whether the p-Cofilin expression is different between GST-Nogo66 addition and GST-Nogo66 plus Pgk1 addition into NSC34 cultured in DM. Compared to the control group in which GST was added in DM, NSC34 cells cultured in GST-Nogo66-added DM exhibited the increase of p-ROCK2-Y256 and p-EGFR-Y1173 expression and the decrease of p-Akt-S473 (Figure 5—figure supplement 3A), suggesting GST-Nogo66 addition did activate NgR response pathway. Interestingly, the increased p-ROCK2-Y256 and p-EGFR-Y1173 and the decreased p-Akt-S473 remained unchanged in NSC34 cultured in GST-Nogo66 with Pgk1 addition (Figure 5—figure supplement 3A), suggesting the addition of Pgk1 did not affect Nogo66/NgR pathway. However, compared to GST-Nogo66-added cells, the GST-Nogo66 plus Pgk1-treated NSC34 cells exhibited the reduction of p-Cpfilin-S3 expression through decreasing p-Limk-S323 but not through p-Limk-T508 (Figure 5—figure supplement 3B), suggesting the reduced expression of p-Cofilin is due to the presence of Pgk1 through pathway other than Nogo66/NgR interaction.

Second, we determined whether the p-Cofilin-S3 expression is still reduced in NSC34 by addition of Pgk1 into NSC34 cultured in DM when NgR receptor is blocked by NAP2, a NgR receptor antagonist peptide. The results showed that addition of NAP2 in culture led to the reduction of ROCK2 expression in NSC34 cells, which was consistent with the result reported by Sun et al. (2016) indicating that NAP2 is able to block NgR receptor (Figure 5—figure supplement 3C). However, unlike the increased ROCK2 in NSC34 cultured in NAP2 addition, the ROCK2 expression remained unchanged in NSC34 cells cultured in NAP2 plus Pgk1 addition (Figure 5—figure supplement 3C). On contrast, the expression of p-Pak1-T423/p-P38-T180/p-MK2-T334/p-Limk1-S323/p-Cofiln-S3 axis was reduced (Figure 5—figure supplement 3D).

Taken together, we suggest that the reduction of p-Cofilin mediated by ePgk1 is not through the Nogo66/NgR interaction in neuronal cells since ePgk1 addition can be still functional to reduce p-Pak1/p-P38/p-MK2/p-Limk1-S323/p-Cofiln axis without the presence of NgR receptor of neuronal cells. In sum, ePgk1 triggers a reduction in p-Cofilin-S3, in turn promoting NOM through decreasing a novel p-Pak1-T423/p-P38-T180/p-MK2-T334/p-Limk1-S323 axis via reducing Rac1-GTP activity in neuronal cells. And, the molecular pathway ePgk1 involved in NOM is independent of the Nogo66/NgR interaction.

Muscle-specific NogoA-overexpression has been shown to disassemble NMJ in animal models, exhibiting muscle atrophy and reduced movement (Jokic et al., 2006; Lin et al., 2019). Noting the colocalization of the presynaptic markers synaptic vesicle glycoprotein 2A (SV2) and Neurofilament-H (NF-H), as well as the postsynaptic marker acetylcholine receptor labeled with α-Bungarotoxin (α-BTX), we asked if ePgk1 could re-establish NMJ integrity after its disassembly by NogoA overexpression. While we observed denervation at NMJ after Rtn4al/NogoA induction in adult zebrafish (Figure 6—figure supplement 1), as evidenced by reduced colocalization of presynaptic and postsynaptic markers, this defect saw much improvement by addition of Pgk1, but not GFP control, through intramuscular injection (Figure 6A–G), suggesting ePgk1 supports NMJ integrity. Although adult zebrafish muscle injected with Pgk1 did exhibit delayed NMJ denervation, it could still retain its normal biological activity for two weeks after a single shot (Figure 6—figure supplement 2).

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We then used an ALS cell line and mouse model to determine any beneficial effects of Pgk1 supplementation. We first differentiated motor neurons derived from human induced pluripotent stem cells (iPSCs) harboring human SOD1 mutant (G85R) (Figure 6—figure supplement 3) and then added Pgk1. Pgk1 addition reduced p-Cofilin through decreasing p-Limk1-S323 (Figure 6—figure supplement 4), as noted above, indicating that ePgk1 induces the same signal transduction pathway as that determined from the mouse cell line and human motor neurons.

Next, to continue exploring this question in vivo, we performed intramuscular injection of Pgk1 into the gastrocnemius muscle of the right hind leg of 60-day-old transgenic SOD1 G93A mice, followed by another injection every 15 days until mice were 120 days old. We quantified the proportion of innervated NMJ in the gastrocnemius muscle of the right hind leg of mice. Compared to the control group, the proportion of innervated NMJ of WT mice injected with PBS (WT/PBS), which was normalized as 100%, the proportion of innervated NMJ of SOD1 G93A/PBS was 13 ± 0.02%, indicating that the signal of motor neuron axon and axon terminal (NF-H/SV2) was significantly reduced. However, the proportion of innervated NMJ of SOD1 G93A mice injected with Pgk1 (SOD1 G93A/Pgk1) was 57 ± 0.08%, suggesting that supplementary addition of Pgk1 could rescue NMJ denervation (Figure 6H–N).

Furthermore, we examined the muscle contraction ability of hind leg in 130-day-old mice. In the WT/PBS group, the muscle contraction of both hind legs was normal, exhibiting strong movement. In contrast, in the SOD1 G93A/PBS group, muscle contraction of both hind legs was extremely poor (Figure 6—videos 1–3; Figure 6—figure supplement 5). Nevertheless, in the SOD1 G93A/Pgk1 group, muscle contraction of the left hind leg was as poor as that of the SOD1 G93A/PBS mice. Interestingly, muscle contraction of Pgk1-injected right hind leg remained functional, exhibiting a superior movement (Figure 6—videos 4–6). Exercise capability value of the SOD1 G93A/Pgk1 group was significantly higher than that of the SOD1 G93A/PBS group (Figure 6—figure supplement 5). Additionally, we found that the proportion of innervated NMJ in the gastrocnemius muscle of the Pgk1-injected right hind leg was increased compared to that of the left hind leg. Taken together, it is suggested that the Pgk1-injected right hind leg of SOD1 G93A mice could maintain some normal motor neurons to innervate muscle contraction.

Our collective findings reveal a novel target against the inhibition of neuron maturation and NOM by overexpression of NogoA through a Nogo66-independent pathway. NogoA has been known to inhibit neuron regeneration via Nogo66 and NiGΔ20 domains. To date, only Nogo66-interacting receptors were found. In this study, we revealed that overexpression of NogoA in muscle inhibits NOM through decreasing Pgk1 secretion (ePgk1), but without inducing the canonical Nogo66/NgR pathway. Here, we demonstrated that the addition of Pgk1 to media could promote NOM by reducing p-Cofilin-S3 through decreasing the phosphorylation of the Pak1/P38/MK2/Limk1-S323 signaling pathway via decreased Rac1-GTP (Figure 7). Whether NogoA also affects the secretion of other trophic factors to inhibit neuronal maturation and maintain NMJ integrity remains a topic for further study.

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The expression of NogoA can generally affect the secretion of proteins. For example, the amount of insulin secreted from beta cells is increased in NogoA-deficient mice (Bonal et al., 2013). In PC12 cells, Zhong et al. (2015) demonstrated that silencing NogoA results in reducing the secretion of inflammatory factors, such as tumor necrosis factor-alpha and interleukin-6 . On the other hand, overexpression of NogoA increases Aβ₄₂ secretion in rat cerebral cortical neurons (Xiao et al., 2012). In this study, when NSC34 motoneuron cells were cultured in CM obtained from cultured Sol8 myoblast cells, which harbor a NogoA-overexpression plasmid, we found that NOM was compromised. Furthermore, we demonstrated that Pgk1 is a key factor contributing to the locomotive properties of neurons in terms of NOM, neurite-bearing cell number and the decrease of p-Cofilin in neuron cells. This finding further demonstrates that improper crosstalk between muscle and nerve tissues impedes the NOM. Although Pgk1 addition could rescue failure NOM induced by culturing NSC34 cells in Sol8-NogoA CM, we noticed that the efficacy of enhancement mediated by Pgk1 alone could barely reach levels as high as those of NSC34 cells cultured in CM from Sol8-vector, suggesting that Pgk1 may not be the only secreted protein involved in enhancing synaptic growth.

To address the reduced secretion of Pgk1 from muscle cells by NogoA, as well as inhibition of the neuronal intracellular Rac1 pathway by ePgk1, we turn to the literature. First, Lay et al. (2000) found that Pgk1 secreted from tumor cells exhibits disulfide reductase activity, resulting in the release of angiostatin through reducing the disulfide bond of plasmin. Wang et al. (2007) also reported that CXCR4 could positively regulate the expression and secretion of Pgk1 in cancer cells. Overexpression of Pgk1 causes an increase of angiostatin, which, in turn, reduces the secretion of vascular endothelial growth factor and interleukin-8. At metastatic sites, a high level of CXCL12 reduces Pgk1 expression, resulting in reduced angiostatin function, which activates angiogenesis. Interestingly, when Rtn4a/NogoA is overexpressed in the somite boundary cells of zebrafish embryos, Lin et al. (2017) found that the expression of Cxcr4a is decreased. Aase et al. (2007) and Yi et al. (2011) reported that angiomotin (Amot), a cell membrane receptor of angiostatin, negatively controls Rac1 activity in endothelial and epithelial cells. Based on this convincing evidence from the literature, we speculate that NogoA/Rtn4a-overexpressed muscle cells may cause a decrease of Cxcr4a expression, resulting in reducing of Pgk1 secretion. The reduced ePgk1 may then cause a decrease in extracellular proteins, such as angiostatin, which, in turn, would increase the activity of Rac1, finally causing a physiological microenvironment for motor neurons that would not favor neurite outgrowth.

Boyd et al. (2017) reported the intracellular, bioenergetic function of Pgk1 in neuron cells, while we described the effect of Pgk1 secreted from muscle cells on NOM through a bioenergetic- and Nogo66-independent pathway. As Boyd et al. (2017) reported, when Pgk1 is insufficiently expressed, its participation in the glycolysis pathway is not effective, resulting in a reduced production of ATP, which, in turn, causes impaired neurological function. Thus, they concluded that Pgk1 is expressed in motor neurons, but also has a bioenergetics function. However, since the effect of pgk1-MO and pgk1 mRNA was not studied in the context of expression in muscle tissue, it remains unclear if the motor neuron phenotype results from the direct effect of Pgk1 within neurons or from indirect effect by defective function of surrounding cells or tissues. Therefore, in this study, we employed a muscle-specific promoter combined with the CRISPR/Cas9 system to specifically knock down Pgk1 in muscle cells, and we found that NOM was inhibited in adjacent nerve cells. Interestingly, when we used the muscle promoter to specifically overexpress Pgk1 in muscle cells, the failed NOM induced by NogoA/Rtn4al overexpression in muscle cells could be rescued. Additionally, we demonstrated that addition of Pgk1 in CM and DM had no impact on the efficiency of intracellular glycolysis in cultured NSC34 cells. In fact, the addition of Pgk1 enables the enhancement of NOM in NSC34 cells, even under reduced glycolytic condition. Additionally, when Pgk1 was ubiquitously knocked down, as Boyd et al. (2017) reported, the total amount of Pgk1 was reduced, including the small amount of secreted Pgk1 from muscle cells, resulting in a phenotype similar to what we observed in this study. Based on this line of evidence, we demonstrated that ePgk1 enhances NOM in a manner functionally independent of its intracellular, canonical role as a supplier of energy, as described by Boyd et al. (2017).

Thus, we have identified a Pgk1/p-Limk1-S323/p-Cofilin-S3 axis in which secreted Pgk1 emerges as an extracellular trigger in the NOM. As illustrated in Figure 6, the following cascade of events takes place. ePgk1 decreases p-Limk1 at S323, which, in turn, results in the decrease of p-Cofilin-S3 and, finally, the rescue of NOM inhibited by culturing in Sol8-NogoA CM. In the process of investigating this novel signaling pathway, we found that overexpression of NogoA in muscle cells resulted in altering the amount of some secreted proteins in muscle cells. For example, the amount of secreted Pgk1 is reduced, which, in turn, inhibits the NOM. Meanwhile, we know that NogoA, as marker of morbidity, overexpresses in the muscle cells of ALS patients at an early stage (Pradat et al., 2007). Because NogoA does not contain a signal peptide (Chen et al., 2000), it is barely detectable in patients’ blood (Harel et al., 2009). Therefore, we have detected a property of Pgk1 in serum that may serve as an indicator of incipient ALS. More recently, Meininger et al. (2017) reported that treatment with Ozanezumab, a humanized monoclonal antibody against NogoA, showed no significant improvement for ALS patients in phase two trial, suggesting that NogoA may not be an effective therapeutic target. Moving away from this target, we proposed in the present study that secreted protein from NogoA-overexpressed muscle cells is a highly promising alternative therapeutic target. More specifically, we showed that the reduced amount of secreted Pgk1 causes neurodegeneration and that the restorative effect of Pgk1 on NOM does not arise through the Nogo66/NgR/ROCK/Limk1-T508 pathway. Although intercellular Nogo66/NgR interaction is blocked by NogoA antibody, it may not change the fact that NogoA is still overexpressed within muscle cells, resulting in the decrease of Pgk1 secretion. Meanwhile, Pgk1 deficiency is inherited as an X-linked recessive genetic trait. It is caused by mutated Pgk1, the folding and stability of which are altered (Chiarelli et al., 2012; Pey et al., 2014). Pgk1 deficiency is accompanied by suffering from muscle lesions, motor neuron defect, neurological dysfunction and myopathy (Valentini et al., 2013; Matsumaru et al., 2017), as well as susceptibility to Parkinson’s disease (Sakaue et al., 2017). Therefore, it can be reasonably speculated that the abnormal secretion of Pgk1 may affect neuronal development, which supports our finding that Pgk1 secreted from muscle cells plays a novel role in NOM and neuronal development.

The lentivirus vectors pAS4.1w.Pbsd-aOn and pAS4.1w.Ppuro-aOn, purchased from the National RNAi Core Facility Platform, Academia Sinica, Taiwan, were used to establish Sol8 myoblasts (RRID:CVCL_6449), which express transgene conditionally. The Sol8-vector was a plasmid pAS4.1w.Ppuro-aOn, while Sol8-NogoA was a plasmid pAS4.1w.Ppuro-aOn with an insert of NogoA cDNA, pAS4.1w.Ppuro-aOn-NogoA. Expression plasmids pCMVΔR8.91 and pMD.G, and lentivirus vector were transfected into cells by Lipofectamine 2000 (Invitrogen) with a 0.9:0.1:1 ratio, respectively, according to the manufacturer’s protocol. The culture medium containing infected virus particles of HEK293T cells (RRID:CVCL_0063, confirmed negative for mycoplasma contamination) was collected at 24 and 48 hr post-transfection after spinning down at 1250 rpm at room temperature. When Sol8 cells grew up to 40% confluency, the supernatant was saved and transferred to Sol8 myoblasts with addition of Polybrene (Sigma) at the concentration of 8 μg/ml for 24 hr. After incubation, the medium was replaced by a fresh growth medium containing Puromycin at the concentration of 4 μg/ml.

After cells reached at 90% confluency, medium was replaced by horse serum (HS)-containing differentiation medium DMEM (2.5% HS, 100 units/ml Penicillin and 0.1 mg Streptomycin) containing Dox at 1 μg/ml. After 24 hr incubation, medium was again replaced by FBS-containing DMEM containing Dox. After another 24 hr incubation, the conditioned medium of Sol8 myoblasts was collected and centrifuged at 1000 rpm. The saved supernatant was used to culture NSC34 cells (Cashman et al., 1992) until they reached at 60% confluency. Medium was then replaced by fresh FBS-containing DMEM containing Dox and continuously cultured for 24 hr. The conditioned media of cultured Sol8 myoblasts was collected by centrifugation and used to culture NSC34 cells for 48 hr. Then, cell lysate was obtained for Western blot analysis.

NSC34 cells were seeded in 10% FBS DMEM medium onto six-well plates at cell density of 3 × 10⁴ per well. After 24 hr incubation, culture medium was replaced by 2.5% FBS DMEM medium to stimulate differentiation of NSC34. After another 24 hr incubation, medium was exchanged for Sol8-conditioned medium and incubated for 96 hr. Finally, cells were fixed with 4% PFA overnight. Neurites extending from each cell were captured by microscopy (Leica) under 200 times magnification and labeled by Neurolucida Neuron Tracing Software 9.0 (Neurolucida 9.0). For each image, 40 cells cultured in either CM or DM were randomly chosen to measure the length of neurites. Data were obtained from three independent trials (120 cells in total). To profile neurite length distribution, we grouped the neurite length into several groups, namely every 25 intervals under 50 µm, every 50 intervals under 200 µm, and 201–300 and 301–600 µm, and calculated the number of cells in each range. To count the number of neurite-bearing cells, we calculated the number of cells with neurites at least 50 μm in length in each range.

The procedures of frozen section and immunostaining of zebrafish and mice tissues and cells were previously described by Lin et al. (2019), except primary antibodies, including anti-Syn1 (RRID:AB_2200097; 1:250), anti-neurofilament H (NF-H) (RRID:AB_91202; 1:500) and anti-acetyl-tubulin (RRID:AB_297928; 1:500), and secondary antibodies, including anti-rabbit Cy2 (RRID:AB_827264; 1:200) and Alexa Fluor 488-conjugated Goat anti-mouse IgG (RRID:AB_2576208; 1:500), were used. The patterns were observed under a Laser Scanning Confocal Fluorescence Microscope (Zeiss). Nuclei were counterstained with 4’,6-diamidino-2-phenylindole (DAPI) (Sigma).

Total proteins of CM from cultured muscle cells in Sol8-NogoA and Sol8-vector were separately collected. Before total proteins were analyzed on 2D SDS-PAGE, Cibacron Blue F3G-A and ammonium sulfate precipitation were performed to remove albumin contained in CM (Colantonio et al., 2005), while remaining proteins were mixed with acetone, precipitated by centrifugation, and dissolved in rehydration buffer (8 M urea, 2% NP-40, and 100 mM DTT). The strips were subjected to rehydration overnight and isoelectric focusing. After isoelectric focusing, the strips were removed and equilibrated. The equilibrated strips were run on the gel as previously described by Sanchez et al. (1999). Gels were stained by Coomassie blue and dried between two sheets of cellophane. Protein spots were excised and subjected to in-gel digestion. The procedures of in-gel digestion and LC/MS-MS analyses were done as previously described in Fu et al. (2012). First, we employed 2D electrophoresis to analyze total protein content in CM. Next, we selected a total of 20 spots at random, all displaying more or less intensity, and we then made a comparison between the Sol8-NogoA CM protein pattern and that of Sol8-vector CM. Following LC MS/MS, 42 candidate proteins were identified (see Figure 1—figure supplement 2). We picked up 10 proteins reduced in Sol8-NogoA CM, cloned their cDNAs, and performed genetic engineering to overexpress these genes individually in Sol8-NogoA cells. We also picked up 10 proteins enhanced in Sol8-NogoA CM, determined their cDNAs, and designed the corresponding siRNAs to knock down their expression individually in Sol8-NogoA cells. All corresponding CMs were collected to culture NSC34 cells separately. After 48 hr in culture, the degree of p-Cofilin expression in NSC34 cells was quantified, and the genes exhibiting lower expression of p-Cofilin were further selected.

Zebrafish (Danio rerio) wild-type AB strain (RRID:ZIRC_ZL1) and transgenic lines Tg(mnx1:GFP) (RRID:ZIRC_ZL1163) (Flanagan-Steet et al., 2005), purchased from ZIRC, and Tg(Zα:TetON-Rtn4al) (Lin et al., 2019) were used. Production and stage identification of embryos followed the description by Lin et al. (2019).

The Mackay Medical College Institutional Animal Care and Use Committee (IACUC) reviewed and approved the protocol described below (MMC-A1060009).

The sgRNAs of pgk1 (TGGACGTGAAAGGAAAGC) and pgam2 (CCTGGAGGAGGCGAAACG) were subcloned into plasmid pDR274 and transcribed into mRNAs in vitro by the mMESSAGE mMACHINE T7 ULTRA kit (Life Technologies). The linearized plasmid pZα-Cas9, in which Cas9 was driven by the muscle-specific alpha-actin promoter of zebrafish, was coinjected separately with pgk1 and pgam2 (control) sgRNAs into zebrafish embryos, and phenotypes were observed at 30 hpf.

The coding region of Limk1 cDNA (NM_010717.3) fused with FLAG-tag was inserted into pCS2⁺ to generate pCS2-Limk1-flag. Mutated forms of Limk1, such as Limk1 S323A and T508V, were obtained by PCR and inserted into pCS2⁺ to generate pCS2-Limk1-S323A-flag and pCS2-Limk1-T508V-flag, respectively. Similarly, Pak1 cDNA (NM_011035.2) with FLAG-tag was inserted into pcDNA3 to generate pcDNA3-Pak1-flag. Additionally, zebrafish pgk1 (NM_213387.1) fused with P2A-RFP and Cas9 fused with P2A-RFP were synthesized and engineered with zebrafish muscle-specific α-actin promoter to generate pZα-Pgk1 and pZα-Cas9 for zebrafish embryo microinjection.

Procedure of the dissociation of zebrafish embryonic cells was followed the previous study reported by Lin et al. (2017) with some modifications. Briefly, the WT embryos and embryos injected with pZα-Cas9, pZα-Cas9 plus pgk1 sgRNA, pZα-Cas9 plus pgam2 sgRNA and pZα-Pgk1 at 30 hpf were incubated with trypsin (Sigma) for 20 min at room temperature. After treatment, embryos were shattered by gently pipetting, followed by separating cells completely from tissues. Then, the RFP-expressing cells were sorted and collected by Cell Sorters (BD FACSAria III).

Protein purification followed the procedures described by Fu et al. (2017). Plasmids expressing FLAG-fusion proteins were transfected into the HEK293T cell line. Transfected cells were lysed using Pierce IP lysis buffer (ThermoFisher Scientific; TFS) with protease inhibitor cocktail (Roche). After cell debris was removed by centrifugation, anti-FLAG M2 affinity gel beads were added to cell extracts and incubated for 16 hr at 4°C. Beads-FLAG-protein complex was eluted by incubation with 3X FLAG peptide for one hr. The FLAG-fusion proteins eluate was restored and used for the following experiment. Plasmids pGEX-GST and pGEX-GST-Nogo66 were used to express recombinant proteins using 0.1 mM Isopropyl β-D-1-thiogalactopyranoside induction for 1 hr at 37°C in an Escherichia coli BL21 (ATCC BAA-1025TM) expression system and purified by Glutathione resin (Clontech).

Total proteins extracted from embryos were analyzed on a 10% SDS-PAGE followed by Western blot analysis according to the procedures described by Lin et al. (2017). The antibodies against NogoA (RRID:AB_650319; 1:500), Cofilin (RRID:AB_10622000; 1:1000), phosphorylated Cofilin at S3 (RRID:AB_2080597; 1:1000), Rho-associated protein kinase 2 (ROCK2) (RRID:AB_10829468; 1:2000), phosphorylated ROCK2 at Y256 (RRID:AB_2182301; 1:2000), Epidermal growth factor receptor (EGFR) (RRID:AB_2246311; 1:500), phosphorylated EGFR at Y1173 (RRID:AB_331795; 1:2000), Akt (Protein kinase B; PKB) (RRID:AB_329827; 1:1000), phosphorylated Akt at S473 (RRID:AB_2315049; 1:1000), LIM domain kinase 1 (Limk1) (RRID:AB_648350; 1:500), phosphorylated Limk1 at S323 (RRID:AB_2136940; 1:1000), phosphorylated Limk1 at T508 (RRID:AB_2136943; 1:500), Phosphoglycerate kinase 1 (Pgk1) (RRID:AB_2161220; 1:2000) (RRID:AB_2268000 for zebrafish; 1:1000), Phosphoglycerate mutase 2 (Pgam2) (RRID:AB_1951200; 1:1000), Ras-related C3 botulinum toxin substrate 1 (Rac1) (RRID:AB_2721173; 1:500), p21-activated kinase 1 (Pak1) (RRID:AB_330222; 1:1000), phosphorylated Pak1 at T423 (RRID:AB_330220; 1:1000), P38 mitogen-activated protein kinases (P38) (RRID:AB_330713; 1:1000), phosphorylated P38 at T180 (RRID:AB_331641; 1:1000), MAP kinase-activated protein kinase 2 (MK2) (RRID:AB_10694238; 1:1000), phosphorylated MK2 at T334 (RRID:AB_490936; 1:1000), Growth Associated Protein 43 (GAP43) (RRID:AB_443303; 1:1000), Choline acetyltransferase (ChAT) (RRID:AB_2244867; 1:1000), Microtubule-associated protein 2 (MAP2) (RRID:AB_2138153; 1:1000), α-tubulin (RRID:AB_477579; 1:5000), Myc (RRID:AB_439680; 1:2000), Flag (RRID:AB_446355; 1:5000), rabbit anti-sheep-HRP (RRID:AB_656968; 1:5000), goat anti-mouse-HRP (RRID:AB_955439; 1:5000) and goat anti-rabbit-HRP (RRID:AB_631746; 1:5000) were used.

Seahorse XF glycolysis stress test kit (Seahorse Bioscience, USA) was used to measure the extracellular acidification rate (ECAR; mpH/min), also called the H⁺ production rate, of NSC34 cells. Briefly, NSC34 cells were seeded in a 10% FBS DMEM medium onto SeaHorse XF 24-well plates at a cell density of 2.5 × 10⁵ per well. After 24 hr incubation, culture medium was replaced by a 2.5% FBS DMEM medium to initiate cell differentiation. Then, NSC34 cells were treated with either Pgk1 (Bio-Techne R and D Systems; BT) at 33 ng/ml or 3PO (SA) at 3 M for two days. When medium was replaced by glucose-free Seahorse 24-well XF Cell Culture, cells were continuously incubated in a non-CO₂ incubator at 37°C for 30 min before assay. To measure the ECAR of the surrounding media, glycolytic flux, such as basal glycolysis, glycolytic capacity and glycolytic reserve, was analyzed by the sequential addition of 10 mM glucose, 1 M Oligomycin and 50 mM 2-deoxyglucose in an XF24 flux analyzer.

The generation of healthy- and ALS-iPS cells from peripheral blood mononuclear cells of hSOD1 G85R was approved under the Institutional Review Board of Hualien Tzu Chi General Hospital, Hualien, Taiwan (IRB-105–131-A). Briefly, iPS cells were established with Cytotune-ips 2.0 Sendai Reprogramming kit (TFS) and cultured in Essential eight medium (TFS) on 1.0% hES qualified Matrigel (Becton Dickinson)-coated cell culture dishes. The cells were passaged per 3–5 days using Accutase (Merck-Millipore), mechanically scraped, and then reseeded at a 1:5 to 1:10 ratio. The culture medium was refreshed daily.

When cell confluence was reached at 80% on the culture dish, cells were treated with Accutase for 2–5 min and scraped. The cell clumps were dissociated into 200–300 mm clusters and transferred to noncoated Petri dishes for 48 hr for embryoid body (EB) formation. The EBs were applied by the modified BiSF neural induction method, according to the methods described previously (Chen et al., 2015). The differentiation medium within the first two days was Essential 6 (TFS). Subsequently, on D2 of differentiation, the cell culture medium was changed to DMEM-F12 supplied with 1% N2 supplement (TFS), 1 mM NEAAs and 2 mM glutamate. Neuron-inducing factors, including 3 µM CHIR99021 (SA), 10 µM SB431542 (SA) and 10 ng/ml recombinant human FGF-2 (R and D Systems) were added on D2 for 2 days. On D4, the neural induction medium was removed, and the neurospheres were cultured in neurobasal medium (TFS) with 1% N2 supplement. Motor neuron patterning factors including 0.5 µM retinoic acid (RA, SA), 0.5 µM purmorphamine and 0.5 µM Smoothened Agonist (SAG, SA) were added from D4 for 5 days, and then reduced dosage of motor neuron patterning factors (0.1 µM RA and 0.25 µM SAG) were added for another 5 days. After complete motor neuron patterning, the cells were dissociated into small clumps by Accutase and seeded on 1% Matrigel (TFS)-coated cell culture dishes for neural maturation. The motor neuron progenitors (MNPs) proliferated in neurobasal medium with 2% B27 supplement (TFS), 10 ng/mL brain-derived neurotrophic factor (BT), 10 ng/ml Glial Cell Line-derived Neurotrophic Factor (BT) and 1 µM Dibutyryl-cAMP (dbcAMP, SA). During the passaging of MNPs with Accutase, providing 10 µM Y27632 (SA) or RevitaCell Supplement (TFS) effectively attenuated dissociation-triggered cell death.

We directly injected Pgk1 (2 μg) into muscle of adult fish from transgenic line Tg(Zα:TetON-Rtn4al), followed by Dox immersion for one week when fish exhibited motorneuron degeneration. The percentage of overlapping signals between SV2 and α-BTX were calculated. We also performed intramuscular injection of Pgk1 into the gastrocnemius muscle of the right hind leg of six 60-day-old (at P60) transgenic SOD1 G93A mice (RRID:IMSR_JAX:002726). Each time, a volume of 80 µl Pgk1 dissolved in PBS solution in a concentration of 375 µg/ml was injected. On three of these frozen longitudinal sections, we performed immunofluorescence staining to quantify the proportion of innervated NMJ in the gastrocnemius muscle of the right hind leg at P75. The remaining three were continuously given Pgk1 injections every 15 days until mice were 120 days old. Six (three for NMJ and three for exercise studies) wild-type mice injected with 80 µl PBS (WT/PBS) into the right leg served as the sham group, and six (three for NMJ and three for exercise studies) SOD1 G93A mice injected with PBS (SOD1 G93A/PBS) served as the negative control group. When the SOD1 G93A/PBS and SOD1 G93A/Pgk1 mice were 130 days old, their movement was recorded by video, and exercise capability of both hind legs was evaluated by quantifying the number of leg contractions within one min. Two legs of each mouse were counted separately. The exercise capability value of each mouse was calculated from the increased fold(s) in the number of contractions between injected right leg versus untreated left leg. Data of each group were averaged from three mouse samples and presented as mean ±S.D.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

This work was supported by Prof. Jin-Chuan Sheu, Dr. Hsiao-Ching Nien and Mr. Spencer Lee, the Liver Disease Prevention and Treatment Research Foundation, Taiwan. We thank TC3 and TC5 Technology Commons, College of Life Science, NTU, for providing the FACS, Microscope and 2D PAGE equipments. We also thank Biomedical Instrumental Center, Mackay Medical College, for providing the SeaHorse XF 24 for Glycolysis Stress Test Assay. We also thank Prof. Hsinyu Lee, College of Life Science, NTU, for providing plasmids pCMVΔR8.91 and pMD.G. We also thank Dr. Neil Cashman, University of Toronto, for providing NSC34 cell line.

Animal experimentation: The Mackay Medical College Institutional Animal Care and Use Committee (IACUC) reviewed and approved the protocol described below (MMC-A1060009).

© 2019, Lin et al.

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