snRPN controls the ability of neurons to regenerate axons

Abstract

Background:

Retinal ganglion cells (RGCs) of mammals lose the ability to regenerate injured axons during postnatal maturation, but little is known about the underlying molecular mechanisms.

Objective:

It remains of particular importance to understand the mechanisms of axonal regeneration to develop new therapeutic approaches for nerve injuries.

Methods:

Retinas from newborn to adult monkeys (Callithrix jacchus) 1 were obtained immediately after death and cultured in vitro. Growths of axons were monitored using microscopy and time-lapse video cinematography. Immunohistochemistry, Western blotting, qRT-PCR, and genomics were performed to characterize molecules associated with axonal regeneration and growth. A genomic screen was performed by using retinal explants versus native and non-regenerative explants obtained from eye cadavers on the day of birth, and hybridizing the mRNA with cross-reacting cDNA on conventional human microarrays. Followed the genomic screen, siRNA experiments were conducted to identify the functional involvement of identified candidates.

Results:

Neuron-specific human ribonucleoprotein N (snRPN) was found to be a potential regulator of impaired axonal regeneration during neuronal maturation in these animals. In particular, up-regulation of snRPN was observed during retinal maturation, coinciding with a decline in regenerative ability. Axon regeneration was reactivated in snRPN-knockout retinal ex vivo explants of adult monkey.

Conclusion:

These results suggest that coordinated snRPN-driven activities within the neuron-specific ribonucleoprotein complex regulate the regenerative ability of RGCs in primates, thereby highlighting a potential new role for snRPN within neurons and the possibility of novel postinjury therapies.

Keywords

Axon regeneration snRPN ganglion cells

1 Introduction

The postnatal maturation of mammalian central nervous system (CNS) neurons, including retinal ganglion cells (RGCs), is characterized by gradual loss of the intrinsic ability to regenerate their axons after injury (Chen, Jhaveri, & Schneider, 1995; Goldberg, Klassen, Hua, & Barres, 2002; Mladinic et al., 2005; J. Nicholls & Saunders, 1996). One possible explanation for this failure of regeneration is thought to be the presence of an inhibitory environment in adults (Doron-Mandel, Fainzilber, & Terenzio, 2015; Schwab, 1990). The developmental and age-related decline in axon growth was demonstrated in (ex vivo cultured) explants from rodent (Chen et al., 1995) and monkey retinas (Rose et al., 2008). Various factors such as cyclic adenosine 3’,5’-monophosphate (cAMP) cAMP response element-binding proteins (CREB) (Cai et al., 2001; Gao et al., 2004; Li et al., 2016), B-cell lymphoma/leukemia2 (Bcl-2) (Chen, Schneider, Martinou, & Tonegawa, 1997; Cho et al., 2005), Krüppel-like-factor-4 (KLF-4) (Benowitz, He, & Goldberg, 2017; Moore et al., 2009), Rho-associated kinase (ROCK) (Lehmann et al., 1999; Liu, Gao, & Wang, 2015), brain derived neurotrophic factor (BDNF) (Zheng et al., 2016) and phosphatase and tensin homologue (PTEN/mTOR) (Park et al., 2008) were identified as regeneration-promoting factors in mice. The involvement of the KLF family in regenerative events in murine RGCs (Moore et al., 2009) indicates that gene clusters with noncoding (nc) RNAs exert regulatory effects on nearby protein-encoding genes.

One of these nuclear clusters is represented by the small nuclear ribonucleoprotein N (snRPN)-ubiquitin E3A imprinting cluster on chromosome 15. Mutations in this gene locus are related to different human hereditary disorders such as Angelman-Syndrome (AS) and Prader–Willi-Syndrome (PWS) (Barlow, 2011; Cheon, 2016; Ferguson-Smith, 2011; Kishore & Stamm, 2006; Manzardo & Butler, 2016; Pauler, Barlow, & Hudson, 2012; Plagge, 2012). The splicesosomal snRPs are major components of the cellular mRNA machinery (Nilsen, 2003; Will & Luhrmann, 2001). The assembly of snRPs is mediated by a multi-protein complex called survival of motor neurons (SMN) complex (Battle et al., 2006; Paushkin, Gubitz, Massenet, & Dreyfuss, 2002). One member of the ribonucleoprotein complex is snRPN, which is specifically expressed in neurons and functions in pre-mRNA processing and alternative splicing. Deletions at the snRPN upstream reading frame protein (snURF)-snRPN locus in mouse models cause PWS-like symptoms (Stefan, Portis, Longnecker, & Nicholls, 2005) with severe phenotypes (Cattanach et al., 1992; Kim et al., 2017; R. D. Nicholls, 1999; Yang et al., 1998). However, despite the findings in the mouse models and the mutations found in the snURF/snRPN gene loci in patients with PWS, a direct role in the pathology of this disease caused by mutations of snRPN gene locus could not be confirmed (Cavaille, 2017; Gallagher, Pils, Albalwi, & Francke, 2002; Meguro et al., 2001; Runte, Varon, Horn, Horsthemke, & Buiting, 2005; Schule et al., 2005; Skryabin et al., 2007), and the question of its possible contribution to other neuronal functions remains open. An association between the snRP complex and regeneration of axons has not yet been reported.

In this investigation of the molecular basis for the loss of axon growth in adult RGCs, we first took advantage of the fact that while axon growth from adult RGCs in culture is marginal to virtually absent, vigorous regeneration of axons occurs from RGCs in retinal explants obtained from newborn New-World monkeys of the Callithrix jacchus species (Rose et al., 2008). The aging-related decline in RGC regenerative ability represents a suitable in vitro model to study the underlying mechanisms triggering this decline. Furthermore, using Callithrix jacchus tissue offers the opportunity to hybridization the monkey mRNA with human Affymetrix-microarrays due to the high genomic homology. This permits to identify potentially regeneration-associated human genes or gene fragments.

This report describes the regulation of members of the snRP complex during retinal development and RGC regeneration. Furthermore, by using small interfering (si) RNA, the functional relevance of one member of the snRP complex to axonal regeneration in adult RGCs could be demonstrated. This is the first study to define a new role for snRPN in neurons and to challenge the search for its use as a therapeutic target toward axonal regeneration.

2 Materials and methods

2.1 Animals and cells

Eyes from marmosets (Callithrix jacchus) aged from 1 day (postnatal day 0; n = 12) to 8 years (n = 8) were obtained from eye cadavers provided by the Institute of Regenerative Medicine, University of Münster, Germany. All animal housing was conducted under the guidelines of ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and under the oversight and approval of the University and of the Governmental Institutional Animal Care and Use Committee (permission numbers 8.87-50.10.46.09.018 for removal of monkey eyes from cadavers and 39.32.7.1/39.32.7.2.1 for breeding of monkeys). Human material was obtained from the university eye clinic Muenster under the tenets of the Declaration of Helsinki.

2.2 Preparation and dissection of the retina

The marmoset retinas were prepared as described before (Rose et al., 2008). The animals were euthanized and their eyes removed and placed into oxygenated Hank’s balanced salt solution (HBSS). After disinfection of the whole eye in 5% iodide solution for 5 min, they were washed in HBSS for 5 min. All of the following preparation steps were conducted under sterile conditions. After removal of the vitreous body, the retina was isolated and carefully spread on a sterile nitrocellulose membrane (pore size, 45μm; Sartorius, Göttingen, Germany) using fine forceps. The membrane was divided into several tapered pieces (Ø 8) and placed with the retinal ganglion cell (RGC) layer on poly-D-lysine- and laminin-coated PetriPERM dishes (Sigma-Aldrich, Munich, Germany) (Fig. 1a). Serum-free S4 medium (fabricated in the laboratory containing DMEM as basis medium, 1% penicillin and streptomycin, and 6.5 g/l glucose) was added to the retina culture (1 ml) and incubated for 96 h in a humidified atmosphere containing 5% CO₂. The number of outgrowing axons was determined after 24, 48, and 72 h in culture using an inverted phase-contrast microscope (×200 magnification; Axiovert 135, Carl Zeiss Jena, Germany) or time-lapse video microscopy (Axiovert 35, Carl Zeiss Jena). In the control condition, retinal samples were placed without a filter in the media, cultured for 3 days without outgrowing axons. For the gene-expression experiments, outgrowing RGCs from retinal explants were cutted with a self-fabricated glass-microknife 72 h after retina explantation, and cultured for additional 24 h to achieve a second regeneration (Fig. 1b).

Fig.1

Procedure for the preparation of the marmoset retina (a) generating outgrowing explants with 1) showing the whole eye, 2) the retina on the filter after preparation, 3) one explant piece after cutting the whole retinal into approximately 8 pieces and 4) outgrowing retinal explant pieces on laminin-coated dishes. (b) cutting of retinal explant (exp) regenerated axons displaying the second axonal regeneration.

2.3 Microarrays

The U133A2 gene chip was prepared according to the manufacturer’s instructions. Briefly, total RNA from the different approaches of C. jacchus retina treatments were transcribed using the GeneChip Two-Cycle cDNA Synthesis kit (Affymetrix), followed by DNA labeling using the GeneChip IVT Labeling kit (Affymetrix). Hybridization was performed using the fragmented and labeled DNA at 45 °C for 16 h. Fluorescence signaling was captured using a GeneChip Scanner 3000 (Affymetrix), and the resulting data files were evaluated using the Expressionist Pro software package (GeneData, Basel, Switzerland). Probes exhibiting different expressions in non-regenerating 3 days in vitro retinas, regenerating retinas, and twice regenerating retinas were identified after global chip quality control and fluorescence gradient correction. Normalized data sets were filtered for sets with a change of at least 1.5-fold up- and downregulated transcripts.

2.4 Immunocytochemistry

A standard protocol was used for immunofluorescence staining of cryosections (Bohm et al., 2012). The antibody rabbit anti-snRPN (dilution 1 : 100, Sigma-Aldrich, HAP003482) was used as primary antibody (diluted in 3% fetal calf serum/PBS as a blocking buffer). A TRITC-conjugated anti-rabbit antibody (dilution 1 : 200, T6778, Sigma-Aldrich,) was used as secondary antibody. Nuclei were counterstained with DAPI.

2.5 Western blotting

The retinal tissues were prepared for SDS-PAGE and semidry blotting, as described previously (Mertsch & Thanos, 2014). Detection was performed using the rabbit anti-snRPN antibody (dilution 1 : 1000, Sigma Aldrich); the secondary antibody was a peroxidase-conjugated goat anti-rabbit antibody (dilution 1 : 20,000, Sigma-Aldrich A9169). GAPDH (dilution 1 : 200,000; Sigma Aldrich) or calnexin (dilution 1 : 20,000, Sigma-Aldrich) were used as a housekeeping proteins for verifying equal protein loading. Results were analysed by lane quantification using densitometry measurement (GelQuantNET software) and loading controls were used for standardization.

2.6 Quantitative real-time PCR

Total RNA was isolated from tissue culture using the GenElute Mammalian Total RNA Miniprep Kit (Sigma-Aldrich). Total mRNA (0.5μg) was transcribed into cDNA using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA). Quantitative real-time PCR measurement was performed using the SYBR-Green method. The following primers were used:

snRPN forward, GCAAGGGATCGCTTACACCT;

snRPN reverse, AGGTACTTGCTGCTGCTGAG.

Data were normalized using the housekeeping gene GAPDH with the following primers: GAPDH forward, TGCACCACCAACTGCTTAGC;

GAPDH reverse, GGCATGGACTGTGGTCATGAG.

The equation fold change = 2^–ΔΔct was applied to calculate relative changes in expression. All measurements were done in duplicates and the experiments were performed at least twice.

2.7 RNA interference experiments

Retinal explants from C. jacchus cadavers were used for RNA interference experiments. Retinas were explanted as described above. Tissue transfection was performed 2 h after retinal culture using the HiPerfect transfection reagent (Qiagen, Hilden, Germany), two different siRNAs directed against the human snRPN sequence (Hs_SNRPN_5 siRNA, SI03132255, Hs_SNRPN_79 siRNA, SI04950946), and a non-silencing negative control (AllStars neg. control siRNA, 1027281; all siRNAs were purchased from Qiagen). For transfection, 625 ng of each siRNA and 40μl of the transfection reagent were used in a total volume of 1 ml of Dulbecco’s modified Eagle’s medium (DMEM) and dispensed dropwise onto the tapered pieces of retina after removing their filter. The explants were re-transfected 24 h after the first transfection using 312.5 ng of siRNA and 20μl of HiPerfect transfection reagent in an additional volume of 500μl of DMEM to reach optimal knockdown efficiency in the tissue. SnRPN knockdown was verified using Western blotting and quantitative real-time PCR 48 h after transfection.

2.8 Statistical analysis

Data are averages of 3 biological repeats +/– standard deviation unless otherwise stated. Data was analysed using Excel software. For testing of statistical significance between two groups, student’s t-tests were performed. p < 0.05 was considered statistically significant.

3 Results

3.1 Confirmation of RGC axonal regeneration in culture

To obtain optimal conditions for axonal regeneration in culture, we first established the explant of retinal pieces as shown in four stages in Fig. 1a. The initial experiments consisted of culturing newborn monkey retinal strips in order to obtain vigorous regeneration of cut axons (Fig. 2a). Lower numbers of axons were observed when the strips were obtained from juvenile, 20-day-old retinas (Fig. 2b), and virtually no axons had grown from explants obtained from adult (3-year-old) retinas (Fig. 2c). In addition, retinas obtained at later stages (up to 12 years of age) also showed nearly non axonal regrowth (data not shown). The formation of a visible “carpet” of regenerated axons from newborn explants (Fig. 3a) permitted the mechanical cutting of these axons (Fig. 3b, c), and subsequently “boosting” of potentially regeneration-associated genes driving the second regeneration of these twice-cut axons. Indeed, cutting of axons that had already regenerated did result in a regrowth of axons starting 30 min after injury (Fig. 3c–f).

Fig.2

Age-dependent decline of marmoset (Callithrix jacchus) retinal ganglion cell axons in tissue cultures. (a) Postnatal day (P)0 marmoset explant after 2 days in culture, with a dense carpet of outgrowing axons. (b) P20 marmoset explant after 2 days in culture, with fewer outgrowing axons. (c) Explant from a 3-year-old marmoset (3y) after 2 days in culture, with no sign of axonal growth.

Fig.3

Cut of regenerated axons in newborn monkey retina explants. (a) Outgrowth of retina explant from a newborn monkey before cutting. (b) Retina explant after cutting the outgrowing axons. (c–f) Timeline of axon re-regeneration up to 4 h after cutting, showing axonal regrowth. In panels c–f the cut line is demarcated by two arrows. Scale bars = 50μm.

3.2 Identification of candidate regulated genes

Candidate genes that are regulated during the process of regeneration were identified by gene-expression profiling of retinal explants from monkeys at postnatal day 0 under different conditions, and comparing the profiles between the native control (Fig. 1a), explantation over 3 days in vitro without regeneration, explantation over 3 days with robust regeneration (Fig. 1a), and explantation over 3 days with robust regeneration plus a second cut in vitro and further culture over 1 day with a second regeneration (Fig. 1b). Since a high genetic homology was expected between the C. jacchus and human genomes, human Affymetrix arrays were used for hybridization with the monkey mRNA. This cross-hybridization revealed some human candidates associated with axon regeneration (Table 1). All candidate genes for which the expression changed by more than 1.5-fold were compared between the three regeneration conditions. Members of the snRP complex were found to be differentially regulated. SnRPF and snRPB were down-regulated in the primary regenerated group compared to native control (Table 2); in contrast, snRPN was up-regulated by 1.98-fold compared to native control. All three members (i.e., snRPF, snRPB, and snRPN) were up-regulated in the non-regenerating groups (ii) compared to the native control (Table 2, Supplement 1). SnRPN and snRPF were further up-regulated in the regenerating-cut group (Table 1), indicating that the second injury to the axons boosted the response of these snRPs. Since snRPN is thought to be neuron-specific, and because it exhibited the most dramatic regulation of 3.16-fold within 24 h after cutting in addition to 1.98-fold after initial regeneration, it emerged as the most interesting candidate gene, and so its role was scrutinized further.

Table 1
Selected genes up-regulated in retinas under the following conditions: non-regenerating after 3 days in vitro, regeneration after axotomy, re-regeneration after re-axotomy and 1 day in vitro

Gene Accession number Gene description Relative change Gene function

snRPN NR_003302 Small nuclear ribonucleoprotein polypeptide N 3.16 RNA splicing, neuron specific

SEC61G NM_014302 Sec61 gamma subunit 2.22 ER transmembrane transport

UBE2MP1 NR_002837 Ubiquitin-conjugating enzyme E2M pseudogene 1 2.00 Protein degradation

POLR2G NM_002696 Polymerase (RNA) II (DNA directed) polypeptide G 1.97 Regulation of RNA-polymerase II

HIST1H1E NM_005321 Histone cluster 1, H1e 1.92 Linker, nucleosome assembly

HMP19 NM_015980 Neuron-specific protein family member 2 1.84 Interacts with RIE3

PSMA5 NM_002790 Proteasome subunit, alpha type, 5 1.83 Peptidase

C12orf51 NM_001109662 Chromosome 12 open reading frame 51 1.78 Protein modification, ubiquitination

SCN2A NM_021007 Sodium channel, voltage-gated, type II, alpha subunit 1.74 Axon potential generation in brain

KIF5A NM_004984 Kinesin family member 5A 1.72 Transport along microtubuli

HLF NM_002126 Hepatic leukemia factor 1.70 PAR family transcription factor

snRPF NM_003895 Small nuclear ribonucleoprotein polypeptide F 1.55 RNA splicing

Gene	Accession number	Gene description	Relative change	Gene function
snRPN	NR_003302	Small nuclear ribonucleoprotein polypeptide N	3.16	RNA splicing, neuron specific
SEC61G	NM_014302	Sec61 gamma subunit	2.22	ER transmembrane transport
UBE2MP1	NR_002837	Ubiquitin-conjugating enzyme E2M pseudogene 1	2.00	Protein degradation
POLR2G	NM_002696	Polymerase (RNA) II (DNA directed) polypeptide G	1.97	Regulation of RNA-polymerase II
HIST1H1E	NM_005321	Histone cluster 1, H1e	1.92	Linker, nucleosome assembly
HMP19	NM_015980	Neuron-specific protein family member 2	1.84	Interacts with RIE3
PSMA5	NM_002790	Proteasome subunit, alpha type, 5	1.83	Peptidase
C12orf51	NM_001109662	Chromosome 12 open reading frame 51	1.78	Protein modification, ubiquitination
SCN2A	NM_021007	Sodium channel, voltage-gated, type II, alpha subunit	1.74	Axon potential generation in brain
KIF5A	NM_004984	Kinesin family member 5A	1.72	Transport along microtubuli
HLF	NM_002126	Hepatic leukemia factor	1.70	PAR family transcription factor
snRPF	NM_003895	Small nuclear ribonucleoprotein polypeptide F	1.55	RNA splicing

Table 2

Regulation of the small nuclear ribonucleoprotein (snRP)-complex members in the newborn regenerating C. jacchus retina: (+), up-regulated; (–), down-regulated

	snRPN relative change	snRPB relative change	snRPF relative change
Regenerating vs native	+1.71	–1.92	–1.69
Regenerating vs explanted	+1.98	–1.52	–1.81
Cut of regenerating	+3.16	Unchanged	+1.55

3.3 Expression of snRPN

We next investigated whether snRPN is developmentally regulated in the monkey retina at various developmental stages and in the control adult human retina. On the day of birth, the RGCs and other retinal interneurons were faintly positive for snRPN (Fig. 4a–c). Immunohistochemical staining of sections at a later juvenile (Fig. 4d–f) and adult (Fig. 4g–i) stages of life revealed that the staining of snRPN increased with age. SnRPN was localized within the nuclei of RGCs. The immunohistochemical data were confirmed by immunoblotting data, with human retinal tissue being used as positive control (Fig. 4j). Densitometric measurement revealed a 137% upregulation of snRPN protein in C. jacchus adult probe compared to the P0 probe (set as 100%) (Fig. 4k). Quantitative real-time measurement confirmed the upregulation of snRPN in later stages of C. jacchus (Fig. 4l). Animals at the age of 4month showed an upregulation of mRNA expression of 167% ±2,4% and animals at the age of 22 month of 179% ±11,38% compared to P0 animals (set as 100%). These data showed that snRPN is expressed in retinal neurons including RGCs, and is regulated during postnatal maturation in the primate retina.

Fig.4

Developmental regulation of small nuclear ribonucleoprotein (snRP)N in the fetal and postnatal marmoset monkey retina. (a–c) Newborn (P0), (d–f) 4 months, (g–i) 22 months. (j) Western blots in the P0 and adult (55 months) monkey retina, with actin as a housekeeping protein for the loading control, and snRPN. (k) densitometric measurement of western blot. (l) quantitative real-time results of snRPN mRNA expression in P0, 4month and 22month marmoset retina showing comparable results to the protein expression. GCL, ganglion cell layer; IPL, inner plexiform layer; INL, inner nuclear layer; OPL, outer plexiform layer; ONL, outer nuclear layer.

The expression profile of snRPN protein in retinal explants was also examined by immunohistochemistry and Western blotting. The expression of snRPN in control explants from newborn retinas was consistent (Fig. 5a–c), and was slightly up-regulated in regenerating explants at day 1 in culture (Fig. 5d–f). These immunohistochemical data were confirmed by Western blotting (Fig. 5g). Densitometric measurement revealed an upregulation of snRPN protein of 886% in P0 retinal explants cultivated for 24 h compared to P0 native (set as 100%). After 72 h in culture, the snRPN expression level decreased to 4%. The direct comparison of snRPN protein expression between P0 and adult native retina showed a 736% upregulation of the protein in the adult tissue. In turn, the protein expression data confirmed the microarray profiling results at the mRNA level and supported the hypothesis that snRPN is regulated during the process of axonal regeneration and response to re-injury.

Fig.5

Expression of snRPN in retinal explants from newborn monkey retinas. (a–c) Control explant immediately after plating onto a culture dish, without cultivation. (d–f) Retinal explant after 1 day in culture. (c) and (f) are merged images of a+b and d+e, respectively. (g) Western blots confirmed the immunohistochemistry findings at 1 and 3 days in vitro (div), as well as the up-regulation between native retina at birth (P0) and in adulthood. (h) densitometric measurement of the western blot.

3.4 Influence of snRPN on RGC axon growth

Since snRPN expression increased with age, as evidenced by both quantitative real-time PCR, immunohistochemistry and Western blotting (Fig. 4) of native retina and retinal explants (Fig. 5), we analysed the functional role of snRPN by assessing if its encoding gene regulates axonal growth of RGCs and determined the effects by knocking down snRPN in retinal tissue explants of adult monkeys in vitro.

Retinal explants from C. jacchus monkeys aged from 10.5 months to 8 years were exposed to siRNA and then cultured for 2 days. The effect of siRNA on snRPN was confirmed with both quantitative real-time PCR and immunoblotting (Fig. 6a,b). SnRPN knockdown resulted in a reduction of snRPN expression in tissue explants to 83.00% ±5.53% (mean±SEM) mRNA expression (scrambled control siRNA set as 100%) for siRNA sequence 1, and a reduction in mRNA to 93.00±9.92% for siRNA sequence 2 (Fig. 6a). This reduction in snRPN expression was verified at the protein level for both siRNA sequences (Fig. 6b). Densitometric measurement showed a downregulation of snRPN to 46.97% protein expression for siRNA 1 and to 40.65% expression for siRNA2 compared to scrambled control set as 100% (Fig. 6c). Lengthy axons grew out after treatment with siRNA compared to scrambled controls (Fig. 6d–f); 17.5±3.0 axons grew out per explant, but none were observed in the controls treated with the nonbinding scrambled control siRNA (Fig. 6g). Thus, downregulation of snRPN expression in the adult retina led to a partial recovery of the regenerative potential in adult RGCs. These data indicate that snRPN is directly involved in the initiation of regenerative axonal growth in adult retinas.

Fig.6

Knockdown experiments with snRPN-targeted small interfering RNA (siRNA) in adult monkey explants showing initiation of axonal regeneration. (a) mRNA level of snRPN after siRNA treatment of the retina explants showing a mRNA reduction in the tissue compared to the control (p < 0.001). (b) Western blotting confirmed the reduction of snRPN protein in siRNA-treated explants showing two different siRNA sequences and calnexin as the housekeeping protein. Data are mean and SEM values from experiments performed in triplicate. (c) densitometric measurement of western blot reveals a reduction of snRPN protein expression of 53.06% for siRNA1 and 59.35% for siRNA2 compared to control (set as 100%). (d-e) photos of retinal explants treated with siRNA or control siRNA (d) with a nonbinding control siRNA, and (d, e) with snRPN siRNA from two different animals. (g) Counts of axons in each experimental setup. Data are mean and SEM values from experiments performed 4 times, with a total number of 32 explants, independently; p < 0,001).

4 Discussion

From the clinical point of view, the strategy used in this study to identify human genes with regeneration-relevant potential appears to be a unique tool to overcome ethical impediments. Although humans diverged from New-World monkeys approximately 33 million years ago (Glazko & Nei, 2003), the overall genomic homology (Tatsumoto et al., 2013) appears to be sufficient to permit hybridization of genes related to axonal regeneration. The common marmoset (C. jacchus) has been successfully used as a nonhuman primate experimental animal for studying Huntington’s disease (Hohjoh et al., 2009) and experimental autoimmune encephalomyelitis (Genain & Hauser, 2001), but to the best of our knowledge, there have been no previous reports of cross-hybridization of functional C. jacchus genes with their human orthologues.

Using the ex vivo cultured marmoset retina as a model of axonal injury and repair, we have established a technique for identifying genes and proteins involved in axonal regeneration. Our cross-hybridization screen with marmoset mRNA and human microarrays revealed a regulation of snRPs and, moreover, enhanced snRPN expression after a second axonal injury that was imposed after a primary regeneration. Importantly, this model is the first to reveal a relationship between snRPs and acute neuronal injury and repair. The confirmation of snRPN regulation during maturation and aging of retinal ganglion cells on protein and mRNA level demonstrates that these genes—and perhaps also other members of the snRP complex—are biologically relevant. Moreover, the finding that snRPN expression is increased during retinal maturation implies a role of this ribonucleoprotein (RP) in the process of aging. Analysis of the role of other members of the snRP complex has the potential to reveal crucial information about the interplay between these core proteins that were also identified as being regulated in the screens.

The finding that the snRP complex regulates axon growth in RGCs of the adult marmoset retina has important implications. Although snRPN has been implicated in human neurological diseases such as PWS (Plagge, 2012), this newly identified function of snRPN in axonal repair advances our knowledge of the molecular regulation of axonal regeneration in aged higher vertebrates. First, snRPN targets relevant to axonal regrowth may include genes and proteins that are selectively expressed in RGCs or may be important for growth-cone assembly and motility. Second, the decline in the intrinsic axon regeneration ability of RGCs with age is paralleled by an increase in the expression of snRPN, and perhaps alterations of other members of the snRP complex, as shown by the pattern of snRPB expression, the regulation of which appears to be inverted compared to snRPN. Thus, manipulation of the genes encoding the snRPs may be a suitable way of increasing the intrinsic regenerative ability of mature RGCs that have been damaged by injury.

Up-regulation of members of the snRP complex was observed in this study, and in particular of snRPN in the probes obtained from double-cut and second regenerating retinal explants. The biogenesis of snRPs follows a complex pathway involving nuclear and cytoplasmic stages (Chari et al., 2008; Janas et al., 2011; Neuenkirchen et al., 2015). The assembly of the Sm protein core onto the Sm-binding site snRNA requires the SMN complex (Battle et al., 2006). Following their transcription by RNA polymerase II in the nucleus, the snRPs encoding RNAs (snRNAs) are exported into the cytoplasm where snRP maturation occurs (Battle et al., 2006). Indeed, our microarray data revealed up-regulation of the polymerase RNAII polypeptide G, which is a regulator of RNA-polymerase II. Further potential members of this highly complex cycle of snRP biogenesis may be seen in the regulation of the gene encoding protein transport protein Sec61 gamma subunit, which has a role in transport across the ER (endoplasmic reticulum) membrane (Greenfield & High, 1999), and the gene encoding kinesin family member 5A (KIF5A), which has a role in transport along microtubules (Goulet et al., 2014). Finally, the gene encoding histone cluster 1, H1e (a histone linker of the nucleosome assembly (Hergeth & Schneider, 2015), is regulated together with genes involved in protein ubiquitination, such as the pseudogene encoding ubiquitin-conjugating enzyme E2M pseudogene 1.

Although the downregulation of snRPN in the ex vivo retinal explants reached only 7–26% knockdown efficiency, which is mainly based on the thickness of the tissue, the high amount of different cell types in the explants and the limitation of siRNA concentration (due to toxicity effects of higher siRNA concentrations), a regeneration of axons in this adult tissue was achieved. This result, together with the microarray data and the mRNA/protein data showed treatment- and age-dependent regulation of snRPN and of different snRP-complex members, which leads us to the hypothesis that other snRP-complex members may also be involved in the axonal regeneration. Indeed, snRPB, snRPF, and snRPD were found to be regulated on the microarray chip as well. The particular roles of these members remain to be examined. In addition, the assemblyosome of the snRPs, the SMN complex (Battle et al., 2006), or Gemins (Charroux et al., 2000) may be involved in the reassembly of snRPs in the injured RGCs. There is evidence that the SMN complex has a role in the transport and local translation of axonal mRNAs (Fallini et al., 2014). In particular, the β-actin encoding axonal mRNA is associated with the SMN complex, indicating its involved in mRNA trafficking to growth cones, thus assigning the complex an additional role beyond that of splicing the snRP assembly (Prescott, Bales, James, Trinkle-Mulcahy, & Sleeman, 2014). Further analyses of these proteins are required to obtain a complete understanding of the function of these proteins within RGCs and their role in axonal regeneration.

In summary we have demonstrated for the first time that snRPN plays a functional role in adult axonal regeneration. Additionally, we have confirmed that screening marmoset genes using human Affymetrix gene chips is a valid method to identify possible candidates for further examinations. Further in vivo work is required to evaluate snRPN and other members of the snRP-complex as therapeutic target for nerve regeneration after optical nerve or retinal injuries.

Footnotes

Acknowledgments

The authors thank Mechthild Wissing and Mechthild Langkamp-Flock for their excellent technical assistance and Magdalena Reis for typing the manuscript. We thank Prof. Dr. Stefan Schlatt for providing the primate material.

Supported by German Research Foundation (DFG) grant (Th 386-22-1 to ST).

The supplementary material is available in the electronic version of this article: .

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