Printer Friendly

MAP kinase cascades regulating axon regeneration in C. elegans.


Mitogen-activated protein kinase (MAPK) signaling cascades are evolutionally conserved in eukaryotes from yeast to humans and play key roles in many diverse physiological processes such as development, growth and proliferation, stress responses, and immunity. A MAPK cascade consists of three classes of protein kinases: MAPK, MAPK kinase (MAPKK) and MAPK kinase kinase (MAPKKK). (1) MAPK is activated by MAPKK, which itself is activated by MAPKKK. This activation cascade can be downregulated by phosphatases, in particular by members of the MAPK phosphatase (MKP) family, which can remove phosphate groups from activated MAPK. (2) The MAPKs are subdivided into three main groups: the extracellular signal-regulated kinases (ERK), the c-Jun N-terminal kinases (JNK) and the p38 kinases. (3) The JNK and p38 MAPKs are activated by environmental stresses such as heat, oxidative stress and ionizing radiation and regulate processes such as stress response, apoptosis and inflammation in mammals (Fig. 1). (4-10) The MKK4 and MKK7 MAPKKs have been shown to activate JNK, and the MKK3 and MKK6 MAPKKs serve as the major activators of the p38 MAPK. The individual MAPKKs are themselves phosphorylated and activated by specific MAPKKKs. Different MKPs display different activities toward the ERK, JNK, and p38 MAPKs.

The ease of genetic manipulation has made the nematode Caenorhabditis elegans an excellent model organism for dissecting the molecular mechanisms underlying numerous processes such as programmed cell death, cell migration and differentiation. A number of JNK and p38 MAPK signaling pathways have been described in C. elegans (Fig. 2). (11-17) Genetic studies demonstrate that the C. elegans JNK pathway, consisting of JKK-1 (a MKK7-type MAPKK) and JNK-1 (a JNK-type MAPK), regulates coordinated movement via type D GABAergic (GABA: [gamma]-aminobutyric acid) motor neurons and also plays a role in synaptic vesicle transport. C. elegans possesses another JNK cascade consisting of MLK-1 (a MLK-type MAPKKK), MEK-1 (a MKK7-type MAPKK) and KGB-1 (a JNK-type MAPK), which is involved in the regulation of the response to heavy-metal stress. One of the C. elegans p38 MAPK pathways, consisting of NSY-1 (an ASK-type MAPKKK), SEK-1 (a MKK3/6-type MAPKK) and PMK-1 (a p38-type MAPK), is involved in innate immunity and the oxidative-stress response. The NSY-1 p38 MAPK pathway is also involved in establishing asymmetric cell fate during neuronal development. The second C. elegans p38 MAPK pathway, consisting of DLK-1 (a DLK-type MAPKKK), MKK-4 (a MLK4-type MAPKK) and PMK-3 (a p38-type MAPK), regulates presynaptic development. Another component of the C. elegans JNK and p38 MAPK pathways is the MKP VHP-1, which negatively regulates these MAPK pathways by dephosphorylating KGB-1, PMK-1 and PMK-3. (18,19)

Recent genetic studies have shown that the MLK-1-MEK-1-KGB-1 JNK and DLK-1-MKK-4-PMK-3 p38 MAPK pathways also regulate axon regeneration in C. elegans. Several studies have demonstrated that the homologues of DLK-1 are required for axon regeneration in both Drosophila melanogaster and mice. (20-23) Similarly, JNK-mediated activation of c-Jun is important for axonal outgrowth of neurons in axotomized rat nodose and dorsal root ganglia (DRG) and mouse DRG. (24-26) These discoveries both validate the use of C. elegans as a model system for axon regeneration and suggest that the core machinery that regulates axon regeneration is conserved from worms to mammals.

C. elegans as a model system for axon regeneration

When an axon is severed, the proximal axonal fragment, which is connected to the cell body, initially retracts and forms a swelling called retraction bulb at its end. If conditions are favorable, the retraction bulb is transformed in a growth cone, a dynamic structure, which extends and guides the regenerating axon towards its target. An axon's ability to regenerate after damage depends on both its intrinsic capacity and the external environment, and is regulated by a balance of regeneration-promoting and -inhibiting factors (Fig. 3). (27) In adult mammals, the regenerative capacity of neurons in the central nervous system (CNS) is limited. In mammals, several myelin-associated factors such as myelin-associated glycoprotein (MAG), oligodendrocyte myelin glycoprotein (OMgp) and Nogo inhibit the regeneration of axons in the CNS. (28-33) On the other hand, injured peripheral neurons activate intrinsic signaling pathways that enable axonal regrowth. When an axon is injured, levels of intracellular calcium ([Ca.sup.2+]) and cyclic adenosine monophosphate (cAMP) are increased. (34,35) This increase in cAMP levels activates protein kinase A (PKA), which activates the axon regeneration-promoting transcription factor cAMP response element binding protein (CREB). (36-38) PKA also contributes to the local modification of the cytoskeleton, which is necessary for the formation and maintenance of the growth cone, a specialized structure necessary to initiate regeneration. (39) Upon axon severance, a long-range retrograde signal is transported from the site of injury to the cell body where various transcription factors are upregulated and subsequently contribute to increased protein synthesis, cell survival and neurite outgrowth. (40) The axonal transport of mRNAs, their stabilization and their local translation provide the structural and regulatory proteins required for axon regeneration in mammals. (41) Manipulation of these processes can improve the chances for successful axon regeneration. Nonetheless, our understanding of the axon-regeneration machinery operating in the adult nervous system of mammals remains limited.

C. elegans is rapidly emerging as a genetic model for probing axon regeneration in the mature nervous system. This animal has a transparent body and it is possible to label specific neurons by cell-specific expression of GFP or other fluorescent proteins. Using a laser device, together with a fluorescent microscope, it is possible to perform laser surgery at the axon level. Moreover, axons of the GABAergic D-type motor neurons of C. elegans can successfully regrow after laser surgery (Fig. 4). (42) Soon after the laser surgery approach was developed, it was complemented by the use of [beta]-spectrin mutants whose axons are fragile and spontaneously break, thereby initiating axon regeneration. (43) These tools enabled researchers to discover novel regulators of axon regeneration by mutagenizing worms and looking for mutants that were unable to regenerate their axons, or alternatively, that regenerate axons better than the wild-type controls. A similar approach is to downregulate specific genes by feeding the worms bacteria expressing a library of siRNAs and looking for positive or negative effects on axon regeneration ability. These genetic screens have been successful in identifying the p38 and JNK MAPK pathways specifically required for axon regeneration. (43,44)

p38 and JNK MAPK pathways are crucial for axon regeneration in C. elegans

An RNAi screen for regulators of axon regeneration identified the MAPKKK DLK-1. (43) It was later demonstrated that activation of the p38 MAPK pathway consisting of DLK-1, MKK-4 and PMK-3 is required for axon regeneration of motor and mechanosensory neurons (Fig. 5). (45) DLK-1 p38 MAPK signaling promotes axon regeneration by activating the MAPKAP kinase MAK-2, which functions to stabilize the mRNA encoding the bZip transcription factor CEBP-1. (45) CEBP-1 in turn promotes axon regeneration, although its targets in this context remain unknown. DLK-1 signaling is also involved in the control of microtubule dynamics during axon regeneration. In mature axons the microtubule cytoskeleton is stably maintained, but after axonal injury it is transformed in a highly dynamic state. This transformation is achieved in two steps: an increase in the number of growing microtubules at the injury site and [DELTA]2 modification of [alpha]-tubulin to support persistent microtubule growth. (46) DLK-1 signaling is involved in the control of both of these phases. First, it decreases the localization of the microtubule depolymerizing kinesin-13 family member KLP-7 in axons by phosphorylating the N-terminus of KLP-7 and thereby reducing its affinity for microtubules. DLK-1 signaling also upregulates the cytosolic carboxypeptidase CCPP-6, which modifies [alpha]-tubulin to the more stable [DELTA]2 form and therefore supports persistent microtubule growth. (46) These effects apparently do not depend on CEBP-1. Although the precise steps by which DLK-1 is activated in axon regeneration remain unknown, a [Ca.sup.2+]-dependent mechanism of activation has been recently described. (47) In its inactive form DLK-1 is a heteromer consisting of a shorter, inactive isoform DLK-1S that binds to and inhibits the longer, active isoform DLK-1L. An increase in [Ca.sup.2+] concentration, which occurs for example after axon injury, dissociates DLK-1S from DLK-1L and allows the formation of DLK-1L homodimers, which are the active form. This spatiotemporal mechanism of DLK-1 activation is suitable for its role in axon regeneration.

An RNAi screen for regulators of axon regeneration also identified another MAPKKK, MLK-1, which is involved in the JNK pathway. (43) A followup study demonstrated that mutation of any of the components of the MLK-1--MEK-1--KGB-1 JNK MAPK pathway significantly impairs axon regeneration (Figs. 5, 6). (44) Conversely, overexpression of MLK-1 improves the frequency and timing of growth cone formation, as well as the rate of successful migration of growth cones to their target sites. This effect was observed also in older animals, which usually are very limited in their regeneration ability. One likely target of JNK MAPK signaling is the transcription factor FOS-1. (17) A loss-of-function mutation in fos-1 significantly reduces the formation of growth cones. (48) FOS-1 is phosphorylated at Thr304 by the KGB-1 JNK and expression of a fos-1 mutant lacking this site (T304A mutant) fails to rescue the defect in axon regeneration of fos-1 mutants. These observations suggest that phosphorylation of FOS-1 on Thr-304 by KGB-1 is important for axon regeneration, although it is still unknown what genes regulated by FOS-1 function to promote axon regeneration.

Several regulators involved in axon regeneration are common to both the p38 and JNK pathways (Fig. 5). For example, DLK-1 and MLK-1 are both targeted for ubiquitin-mediated degradation by the E3 ubiquitin ligase RPM-1. (45) Also, the MAPK phosphatase VHP-1 inactivates both PMK-3 and KGB-1. (18,19) As expected, both rpm-1 and vhp-1 mutants display a MAPK-dependent improvement in axon regeneration. (44,45) However, despite the similarities in their regulation, the downstream effectors of the p38 and JNK pathways are probably largely distinct.

Identification of regulators in the JNK MAPK pathway involved in axon regeneration

The JNK MAPK pathway is negatively regulated by the MAPK phosphatase VHP-1 (Fig. 7). (18,44) The importance of this negative regulation of MAPK signaling is underscored by the fact that vhp-1 mutants are arrested in their development in the early larval stage and almost never reach adulthood.18) Inactivating mutations in mlk-1, mek-1 or kgb-1 suppress this vhp-1 developmental arrest, indicating that the vhp-1 phenotype is caused by hyperactivation of the JNK MAPK pathway. This phenotype has made possible a genome-wide RNAi screen for regulators of the JNK MAPK pathway (Figs. 7, 8). Briefly, we generated animals heterozygous for vhp-1 in which one allele is mutant and the other allele is normal and linked on the same chromosome to a GFP-expressing marker. The animals were fed bacteria that express RNAi from an RNAi library targeting most of the worm genes. We then looked for animals that reached adulthood but did not express GFP. Lack of GFP would indicate that these animals had become homozygous for the vhp-1 mutant allele, and survival would presumably be due to RNAi downregulation of a gene that contributes to JNK MAPK pathway activity. The genes discovered by this screen were termed svh genes, for suppressors of vhp-1 lethality (Fig. 8). The important role that the JNK MAPK cascade plays in axon regeneration suggests that the svh genes might also function as regulators of axon regeneration. The following sections describe the roles that three svh gene products play in axon regeneration: the growth factor SVH-1, its tyrosine kinase receptor SVH-2, and the endocannabinoid hydrolase SVH-3/FAAH-1.

The growth factor SVH-1 and its tyrosine kinase receptor SVH-2 regulate axon regeneration via the JNK MAPK pathway

The first gene identified in the screen for vhp-1 suppressors, svh-1, encodes an extracellular protein possessing a C-type lectin domain, a kringle domain, two LDL receptor-like domains, an N-terminal domain related to plasminogen activation peptide and a serine protease domain (Fig. 9). (49) With the exception of the C-type lectin and LDL receptor-like domains, these protein domains are found also in several mammalian proteins: hepatocyte growth factor (HGF), macrophage stimulating protein (MSP) and plasminogen. This suggests that SVH-1 belongs to the HGF/MSP/plasminogen family. The second gene identified, svh-2, encodes a tyrosine kinase receptor, homologous to the HGF receptor Met and the MSP receptor Ron (Fig. 9). (49) Both svh1 and svh-2 mutants are significantly impaired in their axon regeneration ability. Time-lapse movies indicate that these mutants fail to form stable growth cones, which is a required first step for successful axon regeneration. (49) Conversely, overexpression of either svh-1 or svh-2 induces the formation of enlarged and highly mobile growth cones, markedly improves regeneration outcome, and partially rescues the decline in axon regeneration ability that progresses with age. These results indicate that SVH-1 and SVH-2 are both necessary for axon regeneration and are able to promote regeneration when overexpressed. Together with this, the sequence homology of SVH-1 to HGF and of SVH-2 to the HGF receptor Met suggests that SVH-1-SVH-2 might function as a ligand--receptor pair in axon regeneration (Fig. 9).

Localization of svh-1 expression can be assessed using a reporter gene consisting of the promoter region of the svh-1 gene and the coding sequence of the fluorescent protein Venus. Analyses with this marker indicate that SVH-1 is constitutively expressed in a pair of sensory neurons (ADL) in the head. Deleting the signal sequence from the svh-1 gene generates a secretion-deficient mutant protein that accumulates in the ADL neurons and is not functional in axon regeneration. Expression of svh-1 from either a pan-neuronal promoter or a pharyngeal muscle-specific promoter is sufficient to rescue the svh-1 mutant defect. Furthermore, SVH-1 is not expressed in motor neurons, even after axon injury. These results indicate that SVH-1 functions in axon regeneration in a cell non-autonomous manner. The fact that SVH-1 is constitutively secreted would suggest that it has functions in addition to the regulation of axon regeneration. SVH-1 contains an intact protease domain whose activity is not necessary for axon regeneration and thus its protease activity has other functions. Indeed, SVH-1 acts as a protease required for larval growth.50) Similar studies looking at the expression pattern of a reporter gene driven by the svh-2 promoter reveal that svh-2 is normally not expressed in motor neurons. However, several hours after axotomy svh-2 expression is induced in the injured motor neurons. It was also found that svh-2 can rescue the svh-2 defect only when expressed in the same neurons that undergo axon regeneration. These results are consistent with the idea that SVH-2 functions to promote axon regeneration in a cell-autonomous manner. The injury-dependent expression of SVH-2 probably serves to ensure that the constitutively secreted SVH-1 can initiate SVH-2 signaling in motor neurons only when an axon regeneration response is necessary (Fig. 10).

The mammalian homologues of SVH-2, Met and Ron, form a dimer after ligand binding, and transphosphorylate their respective kinase activation loops on tyrosine residues. (51,52) Several experimental results indicate that SVH-2 functions as a tyrosine kinase in a similar manner. We can create SVH-2 dimers by fusing a dimerization leucine zipper motif (Tpr, translocated promoter region) to the kinase domain and C terminus of SVH-2, thus mimicking ligand-induced dimer formation. When expressed in COS-7 cells and subjected to Western blotting, this hybrid protein Tpr::SVH-2C is recognized by an anti-phosphotyrosine antibody, indicating that it is phosphorylated on tyrosine residues. A catalytically inactive mutant form of Tpr::SVH-2C is not recognized by the anti-phosphotyrosine antibody, suggesting that Tpr::SVH-2C autophosphorylates at tyrosine residues. When expressed in C. elegans, the catalytically inactive form of svh-2 is unable to rescue the axon regeneration defect of svh-2 mutants. This demonstrates that this tyrosine kinase activity is essential for SVH-2 function in axon regeneration. (49)

SVH-2 was identified in a screen designed to discover regulators of the JNK MAPK pathway. Western blotting analysis using COS-7 cells expressing MLK-1 MAPKKK and SVH-2 showed that MLK-1 and SVH-2 can physically interact. Additionally, MLK-1 was found to be tyrosine-phosphorylated in the presence of the wild-type, but not a kinase-dead mutant form of SVH-2, suggesting that SVH-2 directly phosphorylates MLK-1. The most likely site in MLK-1 phosphorylated by SVH-2 is the tyrosine in the sequence NPXY, which creates a docking site for the phospho-tyrosine binding (PTB) domain of SHC-1. SHC-1 is an adaptor protein that connects MLK-1 and its downstream MAPKK MEK-1. Western blotting analysis indicates that SHC-1 binds to MLK-1 only when a kinase-active form of SVH-2 is co-expressed, suggesting that SVH2 phosphorylation is necessary for SHC-1 binding to MLK-1 (Fig. 10). (53)

Taken together, these results support a model in which SVH-1 is constitutively expressed and secreted, while its tyrosine kinase receptor SVH-2 is conditionally expressed in neurons in response to axon injury, whereupon it can be activated by SVH 1. SVH-2 activation induces MLK-1 phosphorylation, leading to enhanced JNK MAPK signaling, which promotes the axon regeneration response (Fig. 10). (49)

Endocannabinoids inhibit axon regeneration by suppressing activation of the JNK MAPK pathway

A third gene, svh-3, was also identified in the screen for regulators of the JNK MAPK pathway. svh-3 is identical to faah-1, which encodes the worm homologue of the fatty acid amide hydrolase, FAAH (Fig. 11). (53) FAAH is an enzyme that hydrolyzes and thereby inactivates several types of endocannabinoids, derivatives of polyunsaturated fatty acids such as arachidonic acid. Endocannabinoids regulate various processes such as synaptic plasticity, pain sensation and memory formation and are also found in worms, where they were discovered to mediate the effects of diet on lifespan. (54,55) Mutation of faah-1 significantly reduces axon regeneration. When faah-1 is knocked down by siRNA, the concentrations of various endocannabinoids increase several-fold, indicating that FAAH-1 is required for maintenance of appropriate low levels of endocannabinoids. (55) This suggested that the defect in axon regeneration observed in faah-1 mutants might be due to the abnormal accumulation of endocannabinoids. Indeed, incubation of wild-type worms with medium containing one of the FAAH-1 substrates arachidonoyl ethanolamide (AEA) strongly inhibits axon regeneration. Several other endocannabinoids, such as eicosapentaenoyl ethanolamide (EPEA), have no effect on axon regeneration, indicating that AEA specifically suppresses axon regeneration.

In mammals the endocannabinoids bind to and activate several transmembrane receptors such as CB1 and CB2. The activated CB1/CB2 receptors in turn activate G proteins, which convey signals downstream. If a similar endocannabinoid signaling mechanism exists in worms, then loss-of-function mutations in any of its components should confer resistance to AEA. It is difficult to test this hypothesis on the receptor level because CB1 and CB2 have no apparent worm homologues, but there is a well-conserved homologue for mammalian Go,, GOA-1. (56,57) Indeed, loss-of-function goa-1 mutants are resistant to the effect of AEA on axon regeneration. Conversely, worms expressing a constitutively active mutant form of goa-1 have a reduced axon regeneration rate even in the absence of AEA. These results suggest that GOA-1 is the downstream effector of AEA. Further analysis of this downstream signaling pathway supports a model in which GOA-1 antagonizes the Gq[alpha] protein EGL-30, which in turn activates the phospholipase EGL-8. EGL-8 generates diacyl-glycerol (DAG), an activator of the protein kinase C (PKC) homologue TPA-1. As expected from this model, mutations in egl-30, egl-8 or tpa-1 result in decreased axon regeneration. The most likely target of TPA-1 in the context of axon regeneration is MLK-1 MAPKKK. Within the activation loop of MLK-1 is a consensus site for PKC phosphorylation (sequence RFS), and the serine within this sequence, Ser-355, is phosphorylated by TPA-1. The link between TPA-1 and MLK-1 is further established by the observation that expression of a phospho-mimetic form of MLK1(S355E) can suppress the regeneration defect of tpa-1 mutants. These results confirm that MLK-1 phosphorylation on Ser-355 is a crucial step for its activation and activity in axon regeneration. In summary, increased signaling from the endocannabinoid pathway leads to suppression of the EGL-30--EGL-8--TPA-1 signaling cascade, which suppresses activation of the MLK-1 JNK MAPK cascade and thereby decreases axon regeneration (Fig. 11).


Results from these studies on SVH-1, SVH-2 and SVH-3/FAAH-1 indicate that the activity of MLK-1 MAPKKK in promoting axon regeneration is regulated by at least two independent mechanisms: it is serine phosphorylated by TPA, which induces its kinase activity, and it is tyrosine phosphorylated by SVH-2, which promotes its interaction with the downstream target MEK-1 MAPKK via the adaptor protein SHC-1 (Fig. 11). Thus, MAPK signaling is regulated at multiple steps, which presumably provides spatiotemporal specificity and ensures responsiveness to the appropriate environmental cues. Negative regulation by endocannabinoids might further restrict the timing of the axon regeneration response to times when conditions are favorable. The synthesis of endocannabinoids increases following several types of injury, such as spinal cord injury and nerve inflammation, and these lipids might therefore function as injury signals. (58,59) The regenerating axon constantly depends on contact with extracellular matrix components both for mechanical support and for guidance towards its target and a significant injury might destroy most of the substrate that the growing cone depends on for proper extension and guidance. In these circumstances, the endocannabinoids might provide an injury signal to restrict the axon regeneration response until the surrounding tissue has recovered sufficiently to be able to support the regenerating axon.

It is still unclear how exactly MAPK signaling regulates axon regeneration but at least some of the regulation involves known transcription factors. The downstream effector of DLK-1 p38 MAPK signaling in axon regeneration is the MAPK-activated kinase 2 MAK-2, which regulates stability of the cebp-1 mRNA encoding a bZip family transcription factor. (45) In addition, one target of JNK MAPK signaling is the transcription factor FOS-1. (17,48) However, it is still unknown what genes are regulated by CEBP-1 and FOS-1 to promote axon regeneration.

MAPK signaling might regulate also axon regeneration via its action on the cytoskeleton and associated regulatory factors. Cytoskeleton reorganization is a major step in the transformation of the inert retraction bulb that forms after axotomy into the highly dynamic growth cone. (39) The formation and maintenance of this complex structure requires precise local control of its building blocks and regulators. MAPK signaling has been implicated in numerous aspects of cytoskeleton regulation. JNK exerts its effects on the cytoskeleton by phosphorylating numerous microtubule-associated proteins (MAPs) such as MAP2, MAP1B, tau and the stathmin family microtubule-destabilizing protein SCG10. (60)-63) Recently it was demonstrated that one major role of DLK-1 signaling in axon regeneration is the control of microtubule dynamics via the kinesin-13 family member KLP-7 and the microtubule-modifying enzyme cytosolic carboxypeptidase CCPP-6. (46) The JNK substrate SCG10 may also be a target of MAPK signaling during axon regeneration, as it is upregulated in rat motor and dorsal root ganglion (DRG) neurons following sciatic nerve crush. (64,65)

Many of the MAPK-associated axon regeneration factors discovered in C. elegans have not yet been verified in other animal systems. However, the conserved central role of MAPK signaling in axon regeneration suggests that at least some of these factors may be relevant for axon regeneration in other animals.

doi: 10.2183/pjab.91.63


(1) English, J., Pearson, G., Wilsbacher, J., Swantek, J., Karandikar, M., Xu, S. and Cobb, M.H. (1999) New insights into the control of MAP kinase pathways. Exp. Cell Res. 253, 255-270.

(2) Camps, M., Nichols, A. and Arkinstall, S. (2000) Dual specificity phosphatases: a gene family for control of MAP kinase function. FASEB J. 14, 6-16.

(3) Morrison, D.K. (2012) MAP kinase pathways. Cold Spring Harb. Perspect. Biol. 4, a011254.

(4) Irie, K., Gotoh, Y., Yashar, B.M., Errede, B., Nishida, E. and Matsumoto, K. (1994) Stimulatory effects of yeast and mammalian 14-3-3 proteins on the Raf protein kinase. Science 265, 1716-1719.

(5) Yamaguchi, K., Shirakabe, K., Shibuya, H., Irie, K., Oishi, I., Ueno, N., Taniguchi, T., Nishida, E. and Matsumoto, K. (1995) Identification of a member of the MAPKKK family as a potential mediator of TGF-O signal transduction. Science 270, 2008-2011.

(6) Shibuya, H., Yamaguchi, K., Shirakabe, K., Tonegawa, A., Gotoh, Y., Ueno, N., Irie, K., Nishida, E. and Matsumoto, K. (1996) TAB1: an activator of the TAK1 MAPKKK in TGF-[beta] signal transduction. Science 272, 1179-1182.

(7) Ichijo, H., Nishida, E., Irie, K., ten Dijke, P., Saitoh, M., Moriguchi, T., Takagi, M., Matsumoto, K., Miyazono, K. and Gotoh, Y. (1997) Induction of apoptosis by ASK1, a mammalian MAPKKK that activates SAPK/JNK and p38 signaling pathways. Science 275, 90-94.

(8) Ninomiya-Tsuji, J., Kishimoto, K., Hiyama, A., Inoue, J., Cao, Z. and Matsumoto, K. (1999) The kinase TAK1 can activate the NIK-I[kappa]B as well as the MAP kinase cascade in the IL-1 signalling pathway. Nature 398, 252-256.

(9) Ishitani, T., Ninomiya-Tsuji, J., Nagai, S., Nishita, M., Meneghini, M., Barker, N., Waterman, M., Bowerman, B., Clevers, H., Shibuya, H. and Matsumoto, K. (1999) The TAK1-NLK-MAPK-related pathway antagonizes signalling between [beta]-catenin and transcription factor TCF. Nature 399, 798-802.

(10) Takaesu, G., Kishida, S., Hiyama, A., Yamaguchi, K., Shibuya, H., Irie, K., Ninomiya-Tsuji, J. and Matsumoto, K. (2000) TAB2, a novel adaptor protein, mediates activation of TAK1 MAPKKK by linking TAK1 to TRAF6 in the IL-1 signal transduction pathway. Mol. Cell 5, 649-658.

(11) Meneghini, M.D., Ishitani, T., Carter, J.C., Hisamoto, N., Ninomiya-Tsuji, J., Thorpe, C.J., Hamill, D.R., Matsumoto, K. and Bowerman, B. (1999) MAP kinase and Wnt pathways converge to downregulate an HMG-domain repressor in Caenorhabditis elegans. Nature 399, 793-797.

(12) Kawasaki, M., Hisamoto, N., Iino, Y., Yamamoto, M., Ninomiya-Tsuji, J. and Matsumoto, K. (1999) A Caenorhabditis elegans JNK signal transduction pathway regulates coordinated movement via type-D GABAergic motor neurons. EMBO J. 18, 3604-3615.

(13) Sagasti, A., Hisamoto, N., Hyodo, J., Tanaka-Hino, M., Matsumoto, K. and Bargmann, C.I. (2001) The CaMKII UNC-43 activates the MAPKKK NSY-1 to execute a lateral signaling decision required for asymmetric olfactory neuron fates. Cell 105, 221-232.

(14) Byrd, D.T., Kawasaki, M., Walcoff, M., Hisamoto, N., Matsumoto, K. and Jin, Y. (2001) UNC-16, a JNK-signaling scaffold protein, regulates vesicle transport in C. elegans. Neuron 32, 787-800.

(15) Kim, D.H., Feinbaum, R., Alloing, G., Emerson, F.E., Garsin, D.A., Inoue, H., Tanaka-Hino, M., Hisamoto, N., Matsumoto, K., Tan, M.-W. and Ausubel, F.M. (2002) A conserved p38 MAP kinase pathway in Caenorhabditis elegans innate immunity. Science 297, 623--626.

(16) Inoue, H., Hisamoto, N., An, J.H., Oliveira, R.P., Nishida, E., Blackwell, T.K. and Matsumoto, K. (2005) The C. elegans p38 MAPK pathway regulates nuclear localization of the transcription factor SKN-1 in oxidative stress response. Genes Dev. 19, 2278-2283.

(17) Hattori, A., Mizuno, T., Akamatsu, M., Hisamoto, N. and Matsumoto, K. (2013) The Caenorhabditis elegans JNK signaling pathway activates expression of stress response genes by derepressing the Fos/HDAC repressor complex. PLoS Genet. 9, e1003315.

(18) Mizuno, T., Hisamoto, N., Terada, T., Kondo, T., Adachi, M., Nishida, E., Kim, D.H., Ausubel, F.M. and Matsumoto, K. (2004) The Caenorhabditis elegans MAPK phosphatase VHP-1 mediates a novel JNK-like signaling pathway in stress response. EMBO J. 23, 2226-2234.

(19) Kim, D.H., Liberati, N.T., Mizuno, T., Inoue, H., Hisamoto, N., Matsumoto, K. and Ausubel, F.M. (2004) Integration of Caenorhabditis elegans MAPK pathways mediating immunity and stress resistance by MeK-1 MAPK kinase and VHP-1 MAPK phosphatase. Proc. Natl. Acad. Sci. U.S.A. 101, 10990-10994.

(20) Valakh, V., Walker, L.J., Skeath, J.B. and DiAntonio, A. (2013) Loss of the spectraplakin short stop activates the DLK injury response pathway in Drosophila. J. Neurosci. 33, 17863-17873.

(21) Xiong, X., Wang, X., Ewanek, R., Bhat, P., DiAntonio, A. and Collins, C.A. (2010) Protein turnover of the Wallenda/DLK kinase regulates a retrograde response to axonal injury. J. Cell Biol. 191, 211-223.

(22) Shin, J.E., Cho, Y., Beirowski, B., Milbrandt, J., Cavalli, V. and DiAntonio, A. (2012) Dual leucine zipper kinase is required for retrograde injury signaling and axonal regeneration. Neuron 74, 1015-1022.

(23) Itoh, A., Horiuchi, M., Bannerman, P., Pleasure, D. and Itoh, T. (2009) Impaired regenerative response of primary sensory neurons in ZPK/DLK genetrap mice. Biochem. Biophys. Res. Commun. 383, 258-262.

(24) Lindwall, C., Dahlin, L., Lundborg, G. and Kanje, M. (2004) Inhibition of c-Jun phosphorylation reduces axonal outgrowth of adult rat nodose ganglia and dorsal root ganglia sensory neurons. Mol. Cell. Neurosci. 27, 267-279.

(25) Kenney, A.M. and Kocsis, J.D. (1998) Peripheral axotomy induces long-term c-Jun amino-terminal kinase-1 activation and activator protein-1 binding activity by c-Jun and junD in adult rat dorsal root ganglia in vivo. J. Neurosci. 18, 1318-1328.

(26) Barnat, M., Enslen, H., Propst, F., Davis, R.J., Soares, S. and Nothias, F. (2010) Distinct roles of c-Jun N-terminal kinase isoforms in neurite initiation and elongation during axonal regeneration. J. Neurosci. 30, 7804-7816.

(27) Chen, Z.-L., Yu, W.-M. and Strickland, S. (2007) Peripheral regeneration. Annu. Rev. Neurosci. 30, 209-233.

(28) McKerracher, L., David, S., Jackson, D.L., Kottis, V., Dunn, R.J. and Braun, P.E. (1994) Identification of myelin-associated glycoprotein as a major myelin-derived inhibitor of neurite growth. Neuron 13, 805-811.

(29) Mukhopadhyay, G., Doherty, P., Walsh, F.S., Crocker, P.R. and Filbin, M.T. (1994) A novel role for myelin-associated glycoprotein as an inhibitor of axonal regeneration. Neuron 13, 757-767.

(30) Wang, K.C., Koprivica, V., Kim, J.A., Sivasankaran, R., Guo, Y., Neve, R.L. and He, Z. (2002) Oligodendrocyte-myelin glycoprotein is a Nogo receptor ligand that inhibits neurite outgrowth. Nature 417, 941-944.

(31) Prinjha, R., Moore, S.E., Vinson, M., Blake, S., Morrow, R., Christie, G., Michalovich, D., Simmons, D.L. and Walsh, F.S. (2000) Inhibitor of neurite outgrowth in humans. Nature 403, 383-384.

(32) Chen, M.S., Huber, A.B., van der Haar, M.E., Frank, M., Schnell, L., Spillmann, A.A., Christ, F. and Schwab, M.E. (2000) Nogo-A is a myelin-associated neurite outgrowth inhibitor and an antigen for monoclonal antibody IN-1. Nature 403, 434439.

(33) GrandPre, T., Nakamura, F., Vartanian, T. and Strittmatter, S.M. (2000) Identification of the Nogo inhibitor of axon regeneration as a Reticulon protein. Nature 403, 439-444.

(34) Kulbatski, I., Cook, D.J. and Tator, C.H. (2004) Calcium entry through L-type calcium channels is essential for neurite regeneration in cultured sympathetic neurons. J. Neurotrauma 21, 357374.

(35) Ziv, N.E. and Spira, M.E. (1995) Axotomy induces a transient and localized elevation of the free intracellular calcium concentration to the millimolar range. J. Neurophysiol. 74, 2625-2637.

(36) Appenzeller, O. and Palmer, G. (1972) The cyclic AMP (adenosine 3',5'-phosphate) content of sciatic nerve: changes after nerve crush. Brain Res. 42, 521-524.

(37) Carlsen, R.C. (1982) Axonal transport of adenylate cyclase activity in normal and axotomized frog sciatic nerve. Brain Res. 232, 413-424.

(38) Gao, Y., Deng, K., Hou, J., Bryson, J.B., Barco, A., Nikulina, E., Spencer, T., Mellado, W., Kandel, E.R. and Filbin, M.T. (2004) Activated CREB is sufficient to overcome inhibitors in myelin and promote spinal axon regeneration in vivo. Neuron 44, 609-621.

(39) Hur, E.-M., Saijilafu and Zhou, F.-Q. (2012) Grow ing the growth cone: remodeling the cytoskeleton to promote axon regeneration. Trends Neurosci. 35, 164-174.

(40) Abe, N. and Cavalli, V. (2008) Nerve injury signal ing. Curr. Opin. Neurobiol. 18, 276-283.

(41) Bradke, F., Fawcett, J.W. and Spira, M.E. (2012) Assembly of a new growth cone after axotomy: the precursor to axon regeneration. Nat. Rev. Neurosci. 13, 183-193.

(42) Yanik, M.F., Cinar, H., Cinar, H.N., Chisholm, A.D., Jin, Y. and Ben-Yakar, A. (2004) Neurosurgery: functional regeneration after laser axotomy. Nature 432, 822.

(43) Hammarlund, M., Nix, P., Hauth, L., Jorgensen, E.M. and Bastiani, M. (2009) Axon regeneration requires a conserved MAP kinase pathway. Science 323, 802-806.

(44) Nix, P., Hisamoto, N., Matsumoto, K. and Bastiani, M. (2011) Axon regeneration requires coordinate activation of p38 and JNK MAPK pathways. Proc. Natl. Acad. Sci. U.S.A. 108, 10738-10743.

(45) Yan, D., Wu, Z., Chisholm, A.D. and Jin, Y. (2009) The DLK-1 kinase promotes mRNA stability and local translation in C. elegans synapses and axon regeneration. Cell 138, 1005-1018.

(46) Ghosh-Roy, A., Goncharov, A., Jin, Y. and Chisholm, A.D. (2012) Kinesin-13 and tubulin post-translational modifications regulate microtubule growth in axon regeneration. Dev. Cell 23, 716-728.

(47) Yan, D. and Jin, Y. (2012) Regulation of DLK-1 kinase activity by calcium-mediated dissociation from an inhibitory isoform. Neuron 76, 534-548.

(48) Nix, P., Hammarlund, M., Hauth, L., Lachnit, M., Jorgensen, E.M. and Bastiani, M. (2014) Axon regeneration genes identified by RNAi screening in C. elegans. J. Neurosci. 34, 629-645.

(49) Li, C., Hisamoto, N., Nix, P., Kanao, S., Mizuno, T., Bastiani, M. and Matsumoto, K. (2012) The growth factor SVH-1 regulates axon regeneration in C. elegans via the JNK MAPK cascade. Nat. Neurosci. 15, 551-557.

(50) Hisamoto, N., Li, C., Yoshida, M. and Matsumoto, K. (2014) The C. elegans HGF/plasminogenlike protein SVH-1 has protease-dependent and -independent functions. Cell Rep. 9, 1628-1634.

(51) Ferracini, R., Longati, P., Naldini, L., Vigna, E. and Comoglio, P.M. (1991) Identification of the major autophosphorylation site of the Met/hepatocyte growth factor receptor tyrosine kinase. J. Biol. Chem. 266, 19558-19564.

(52) Gaudino, G., Follenzi, A., Naldini, L., Collesi, C., Santoro, M., Gallo, K.A., Godowski, P.J. and Comoglio, P.M. (1994) RON is a heterodimeric tyrosine kinase receptor activated by the HGF homologue MSP. EMBO J. 13, 3524-3532.

(53) Pastuhov, S.I., Fujiki, K., Nix, P., Kanao, S., Bastiani, M., Matsumoto, K. and Hisamoto, N. (2012) Endocannabinoid-Goa signalling inhibits axon regeneration in Caenorhabditis elegans by antagonizing Gq[alpha]-PKC-JNK signalling. Nat. Commun. 3, e1136.

(54) Di Marzo, V., Bifulco, M. and De Petrocellis, L. (2004) The endocannabinoid system and its therapeutic exploitation. Nat. Rev. Drug Discov. 3, 771-784.

(55) Lucanic, M., Held, J.M., Vantipalli, M.C., Klang, I.M., Graham, J.B., Gibson, B.W., Lithgow, G.J. and Gill, M.S. (2011) N-acylethanolamine signalling mediates the effect of diet on lifespan in Caenorhabditis elegans. Nature 473, 226-229.

(56) Mendel, J.E., Korswagen, H.C., Liu, K.S., Hajdu Cronin, Y.M., Simon, M.I., Plasterk, R.H. and Sternberg, P.W. (1995) Participation of the protein Go in multiple aspects of behavior in C. elegans. Science 267, 1652-1655.

(57) Segalat, L., Elkes, D.A. and Kaplan, J.M. (1995) Modulation of serotonin-controlled behaviors by Go in Caenorhabditis elegans. Science 267, 1648-1651.

(58) Calignano, A., La Rana, G., Giuffrida, A. and Piomelli, D. (1998) Control of pain initiation by endogenous cannabinoids. Nature 394, 277-281.

(59) Eljaschewitsch, E., Witting, A., Mawrin, C., Lee, T., Schmidt, P.M., Wolf, S., Hoertnagl, H., Raine, C.S., Schneider-Stock, R., Nitsch, R. and Ullrich, O. (2006) The endocannabinoid anandamide protects neurons during CNS inflammation by induction of MKP-1 in microglial cells. Neuron 49, 67-79.

(60) Chang, L., Jones, Y., Ellisman, M.H., Goldstein, L.S.B. and Karin, M. (2003) JNK1 is required for maintenance of neuronal microtubules and controls phosphorylation of microtubule-associated proteins. Dev. Cell 4, 521-533.

(61) Goedert, M., Hasegawa, M., Jakes, R., Lawler, S., Cuenda, A. and Cohen, P. (1997) Phosphorylation of microtubule-associated protein tau by stress-activated protein kinases. FEBS Lett. 409, 57-62.

(62) Reynolds, C.H., Utton, M.A., Gibb, G.M., Yates, A. and Anderton, B.H. (1997) Stress-activated protein kinase/c-jun N-terminal kinase phosphorylates tau protein. J. Neurochem. 68, 1736-1744.

(63) Tararuk, T., Ostman, N., Li, W., Bjorkblom, B., Padzik, A., Zdrojewska, J., Hongisto, V., Herdegen, T., Konopka, W., Courtney, M.J. and Coffey, E.T. (2006) JNK1 phosphorylation of SCG10 determines microtubule dynamics and axodendritic length. J. Cell Biol. 173, 265-277.

(64) Mason, M.R.J., Lieberman, A.R., Grenningloh, G. and Anderson, P.N. (2002) Transcriptional upregulation of SCG10 and CAP-23 is correlated with regeneration of the axons of peripheral and central neurons in vivo. Mol. Cell. Neurosci. 20, 595-615.

(65) Voria, I., Hauser, J., Axis, A., Schenker, M., Bichet, S., Kuntzer, T., Grenningloh, G. and BarakatWalter, I. (2006) Improved sciatic nerve regeneration by local thyroid hormone treatment in adult rat is accompanied by increased expression of SCG10. Exp. Neurol. 197, 258-267.

(Received Dec. 24, 2014; accepted Jan. 13, 2015)

By Strahil Iv. PASTUHOV, * [1] Naoki HISAMOTO * [1] and Kunihiro MATSUMOTO * (1), ([dagger])

(Communicated by Shigekazu NAGATA, M.J.A.)

* [1] Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya, Japan.

([dagger]) Correspondence should be addressed: K. Matsumoto, Division of Biological Science, Graduate School of Science, Nagoya University, Chikusa-ku, Nagoya 464-8602, Japan (e-mail:


Kunihiro Matsumoto was born in Nagasaki in 1951, and graduated Faculty of Engineering, Osaka University in 1974. At the second year of his Ph.D. course, he started his research career with studies on signal transduction in yeast as Assistant Professor at Tottori University in 1977. He received his Ph.D. from Osaka University in 1982. He then moved to DNAX Research Institute in California as Senior Scientist in 1985. He continued his studies on signal transduction, especially MAP kinase pathways, in yeast. In 1990, he returned to Japan as Professor at Nagoya University, where he switched his research area to MAP kinase pathways in mammalian cells and the nematode Caenorhabditis elegans. For his great achievements on molecular genetics, he was honored to receive Japanese Genetic Society Young Investigator Award in 1985, Nissan Science Foundation Award in 2001, Kihara Memorial Foundation Award in 2001, Inoue Prize for Science in 2002, Mochida Memorial Foundation Award in 2012, National Prize of Purple Ribbon Medal (Japan) in 2012 and Chunichi Cultural Award in 2013. He becomes the Dean of Graduate School of Science at Nagoya University in 2015.
COPYRIGHT 2015 The Japan Academy
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2015 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Title Annotation:Mitogen-activated protein
Author:Pastuhov, Strahil Iv.; Hisamoto, Naoki; Matsumoto, Kunihiro
Publication:Japan Academy Proceedings Series B: Physical and Biological Sciences
Article Type:Report
Geographic Code:9JAPA
Date:Mar 1, 2015
Previous Article:Toward the detection of gravitational waves under non-Gaussian noises I. locally optimal statistic.
Next Article:Intraovarian control of selective follicular growth and induction of oocyte maturation in mammals.

Terms of use | Privacy policy | Copyright © 2020 Farlex, Inc. | Feedback | For webmasters