Prion protein cleavage fragments regulate adult neural stem cell quiescence through redox modulation of mitochondrial fission and SOD2 expression
Neurogenesis continues in the post-developmental brain throughout life. The ability to stimulate the production of new neurones requires both quiescent and actively proliferating pools of neural stem cells (NSCs). Actively proliferating NSCs ensure that neurogenic demand can be met, whilst the quiescent pool makes certain NSC reserves do not become depleted. The processes preserving the NSC quiescent pool are only just beginning to be defined. Herein, we identify a switch between NSC proliferation and quiescence through changing intracellular redox signalling. We show that N-terminal post-translational cleavage products of the prion protein (PrP) induce a quiescent state, halting NSC cellular growth, migration, and neurite outgrowth. Quiescence is initiated by the PrP cleavage products through reducing intracellular levels of reactive oxygen species. First, inhibition of redox signalling results in increased mitochondrial fission, which rapidly signals quiescence. Thereafter, quiescence is maintained through downstream increases in the expression and activity of superoxide dismutase-2 that reduces mitochondrial superoxide. We further observe that PrP is predominantly cleaved in quiescent NSCs indicating a homeostatic role for this cascade. Our findings provide new insight into the regulation of NSC quiescence, which potentially could influence brain health throughout adult life.
KeywordsNADPH oxidase Nox2 DRP1 Superoxide dismutase SOD2 Reactive oxygen species Mitochondria N1 N2
It is now firmly established that the adult brain contains cells that demonstrate ‘stemness’, i.e., are capable of self-renewal and formation of new brain cells (reviewed in ). Neural stem cells (NSCs) have been detected in human brain tissue from donors up to the age of 72 years  and markers of neurogenesis detected into the ninth decade , indicating the likely importance of these cells throughout adult life. The adult brain contains different populations of NSCs. Type 1 NSCs are quiescent and can be stimulated to become type 2 NSCs, which are actively replicating. Type 2 cells in turn progress to be type 3 migratory cells or neuroblasts [1, 4]. NSC growth and differentiation have been linked with hippocampal learning and memory as well as forgetting [5, 6, 7, 8, 9], and changes in neurogenesis correlate with various mental health issues and neurodegenerative diseases [3, 10].
Neurogenesis is modulated by several neurodegenerative disease-associated proteins or peptides including the prion protein (PrP) [11, 12], which is most widely recognised for its causative role in transmissible neurodegenerative diseases of humans and animals . Research into the role of PrP in neurogenesis has found that PrP expression is linked with enhanced NSC proliferative capacity , associated with increased cell cycling at the expense of differentiation . In addition, PrP has been shown to be part of a receptor complex for soluble neurotoxic Alzheimer’s disease-associated amyloid-beta peptides [15, 16] and the presence or absence of PrP changes NSC self-renewal in response to amyloid-beta peptides , thereby suggesting that a putative neurogenic function of PrP may become corrupted during neurodegenerative disease.
Redox balance has a regulatory role in NSC growth and differentiation [34, 35]. Adult NSCs are not dispersed evenly through the brain but situated in specialised regions, referred to as neurogenic niches . In the neurogenic niche, the environment is highly important for the preservation of the NSC populations and is maintained as a low ROS environment . Increased NSC ROS are associated with stimulation of increased growth, possibly at the expense of depleting the quiescent pool in favour of the actively proliferating cells [34, 35, 38]. In neurodegenerative diseases, such as prion diseases and Alzheimer’s disease, where redox balance is compromised resulting in increased ROS, increased markers of neurogenesis are detected in brain tissue . We hypothesized that the ability of the N-terminal PrP peptides, N1 and N2, to modulate cellular ROS levels could exert significant regulatory effects on NSC growth and maintenance of the quiescent NSC pool.
Prion N-terminal peptides
The prion N-terminal peptides were generated by microwave synthesis and quality controlled by HPLC and mass spectrometry as previously described in Karas et al. . Peptides were also purchased from China Peptides (China).
Adult neural stem cell extraction
NSCs were harvested from the brains of three 8-week-old PrP gene (prnp)−/− (PrP knock-out), C57/B6 wild-type, and Tga20 (PrP over-expressing by approximately eightfold) mice, and transferred into suspension culture as described previously in Haigh et al. . All animal procedures were performed in accordance with University of Melbourne animal ethics committee approval (ethics ID: 1413198.1), operating under the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes.
Routine NSC culture has been described previously . In brief, cells were grown as neurospheres in complete proliferation medium (Stem Cell Technologies, VIC, AUS), supplemented with final concentrations of 10 ng/ml fibroblast growth factor (FGF) and 20 ng/ml epidermal growth factor (EGF) cytokines, and 2 μg/ml heparin (Stem Cell Technologies). Cultures were maintained in a 5% CO2 humidified incubator.
Cells were mixed 1:1 (v/v) with Trypan Blue reagent (Life Technologies, VIC, AUS) and counted using a Countess cell counter (Life Technologies).
Neural colony forming assay (NCFA)
The colony forming assay is described in Collins et al. . Briefly, neurospheres are dissociated into single-cell suspension and seeded in a semi-solid gel matrix made with a 2:1 solution of proliferation medium and collagen (Sigma-Aldrich, VIC, AUS) in 24-well plates at a density of 50,000 cells/ml (25,000 cells/well). Wells were supplemented with 80 μl of fresh proliferation medium every 5 days. Treatments were included at the start of the assay only. Colonies were counted and diameters measured at 21 days post the start of the assay. All colonies at all depths through the matrix were calculated and measured for a minimum of 5 fields or 50 colonies (for conditions where few colonies were found per field of view).
Neurospheres of 80–100 μm diameter were seeded into wells pre-coated with 5 μg/ml poly-d lysine (PDL; Sigma-Aldrich) in H2O solution (for a minimum of 1 h at room temperature) and grown in complete proliferation medium. After 3 days, migration from the central neurosphere was calculated by application of contour ROIs around the surface area into which the cells had spread using NIS Elements 3.0 (Nikon) software package. The area covered after 7 days could not be calculated as migration had proceeded beyond the imaging area.
Neurite outgrowth assay
One hundred thousand cells/well were cultured in complete proliferation medium supplemented with 20 ng/ml FGF for 24 h before transferral into complete differentiation medium (Stem Cell Technologies). The neurite outgrowth assay (Millipore, Thermo-Fisher, VIC, AUS) was carried out as per the manufacturer’s instructions. Well inserts (1 µm pore size) were coated on their under-side with 10 µg/ml laminin in H2O solution for 2 h at 37 °C before NSCs were transferred into the insert. Cells were cultured for 2 days in differentiation medium before assaying outgrowth. To assay neurite extension onto the laminin coated under-side, well inserts were first washed in PBS, then fixed in methanol for 20 min at room temperature, washed again by transferring the insert into PBS, and then stained with neurite stain solution for 30 min. Inserts were washed again in PBS, excess dye removed from the inside of the insert, and cell somas removed by swabbing the inside of the insert. The neurites extending over the under-side of the well insert were then solubilised in extraction buffer and the absorbance read in a Spectrostar (BMG) at 562 nm. Neurite outgrowth, indicated by absorbance of the neurite stain as compared with extraction buffer alone, was calculated using MARS software (BMG).
The standard immunofluorescence protocol used has been described previously [24, 41]. Briefly, cells were fixed in 4% (v/v) paraformaldehyde for 30 min, followed by permeabilization in 0.1% triton-X-100, before blocking in 10% (v/v) FBS, 0.1% (w/v) BSA in PBS for 30 min. Primary and secondary antibody incubations were carried out in 1% (v/v) FBS, 0.1% (w/v) BSA in PBS blocking buffer, with details and relevant concentrations listed in Sup Table 1.
Confocal images were collected using a Leica SP8 (Leica Microsystems, NSW, AUS) and wide-field fluorescence or bright-field images captured using a Nikon Eclipse TE2000-E epi-fluorescence microscope with a Roper Scientific CCD camera (Nikon, Coherent Scientific, SA, AUS). Image analysis was performed using Fiji imaging software  or NIS Elements 3.0 (Nikon). Mitochondrial parameters (minimum of six fields from three independent experiments) were calculated using the mito-morphology macro for image J created by Ruben K. Dagda at the University of Pittsburgh. Image enhancements, despeckling, were performed for inset digital zooms to increase picture clarity.
Pharmacological inhibition of pathway activity
Diphenyleneiodonium chloride (DPI; Sigma-Aldrich) stocks were prepared in water and stored at room temperature. N-[2-(p-Bromocinnamylamino)ethyl]-5-isoquinolinesulfonamide dihydrochloride (H89; Abcam, VIC, AUS) stocks were prepared in water and stored at -20°C.
Cells were seeded at a density of 3.6 × 104 cells/well in 5 μg/ml PDL in H2O solution-coated 96-well plates and allowed to adhere under normal incubator conditions for 24 h before assay.
[3-(4,5-Dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS) metabolism assay
Five microlitres of one-solution MTS reagent (Promega, VIC, AUS) was added per 100 μl of media in each well. Plates were incubated under normal incubator conditions for the duration of the assay. Developed colour was measured at 492 nm in a FluoSTAR/PolarSTAR Optima (BMG Labtech, VIC, AUS).
Lactate dehydrogenase (LDH) cell death assay
Cellular and extracellular LDH was measured using a Cytotoxicity Detection kit (LDH; Roche, VIC, AUS) and cytotoxicity calculated as per the manufacturer’s instructions.
Caspase 3/7 assay
Caspase 3/7 staining and 7-amino-actinomycin D (7-AAD) staining were carried out and quantified using a Muse™ Caspase-3/7 Kit (Roche-Sigma, USA) and a Muse™ Cell Analyzer (Roche-Sigma) with software version 1.4. The cells were loaded as per the manufacturer’s instruction with 5 µl of Caspase-3/7 working solution added to 50 µl cells (diluted to approximately 1 × 106 cells/ml). Cells were incubated for 30 min at 37 °C before addition of 150 µl of 7-ADD working solution and then analysed for apoptotic (but not dead) and dead cells.
Ki67 was assayed using the Muse® Ki67 Proliferation Kit (Roche-Sigma) and a Muse™ Cell Analyzer (Roche-Sigma). Cells were fixed and stained as per the manufacturer’s instructions and the negative control staining (shown in grey shading on the plots) was used to gate the background unstained cell population for analysis using the Muse v1.4 software.
Adenosine triphosphate (ATP) assay
Cellular ATP content was measured by luminescence using Life Sciences ATP assay (Life Technologies) as per the product protocol and normalised for total protein levels determined by BCA assay (Pierce, Thermo-Fisher).
Dichlorofluorescin diacetate (DCFDA) assay
The DCFDA assay for intracellular ROS production has been described previously . Briefly, cells were loaded with CM-H2-DCFDA reagent (Invitrogen) by incubating cells with a 5 µM probe solution in PBS for 20 min at 37 °C. Basal fluorescence was then read to provide a well-background control and test conditions added to start the assay. Fluorescence intensity was monitored for 12 h.
Nicotinamide adenine dinucleotide phosphate (NADPH) consumption assay
NADPH (Arbor Assays, Bio Scientific, VIC, AUS) was diluted to 250 mM stock in sterile water, and aliquoted and stored at – 20 °C until use. At the start of the assay, 1 μM peptide was included in the culture media; then, 25 mM NADPH was added to each well and decay of the NADPH absorbance monitored at 340 nm every 60 s for 10 min and gradients calculated to show consumption of substrate.
Superoxide dismutase (SOD) activity assay
Total SOD activity of cell lysates was determined using a WST-1-based competitive inhibition assay (Abcam), as per the manufacturer’s instructions, with normalisation of activity to total protein as determined by BCA assay.
Peptide:N-glycosidase F (PNGase-F) digests
Pre-denatured lysates were digested using 1 µl PNGase-F per 20 µl protein sample with overnight incubation at 37 °C.
Western blotting was carried out using the Invitrogen NuPAGE/Bolt gel system (Life Technologies) with Criterion (BioRad, VIC, AUS) or iBlot (Invitrogen) transfer and developed as described previously [41, 43]. Coomassie membrane staining was performed as previously described . Total protein staining was used to monitor gel loading/transfer efficiency, because the housekeeping proteins often used as loading controls are involved in the cellular processes being investigated and are, therefore, considered unreliable. Dot blotting was performed by dotting 2 μl of whole-cell lysate onto nitrocellulose membranes. Membranes were blocked and blotted as for standard western blotting procedure. Antibody information is shown in Sup Table 1. Percentage change was calculated by first normalising band signal to total protein and then applying the following equation: % change = (test band signal intensity/control band signal intensity) × 100.
Senescence staining and quantification
Senescence was determined using a β-galactosidase staining kit (Cell Signaling Technologies, Sapphire Biosciences, VIC, AUS), following the manufacturer’s instructions and the blue product solubilised in DMSO by agitated heating as described in Haigh et al. .
Redox-sensor Red ROS assay
Redox-sensor Red (PF-H2TMRos; Life Technologies) at a final concentration of 5 μM in Opti-MEM I reduced serum culture medium (phenol red-free; Invitrogen) was loaded into cells for 10 min and cells were imaged in fresh medium as described previously .
MitoSOX fluorescent indicator probe (Life Technologies) was loaded into cells at a final concentration of 5 μM in normal media for 10 min under standard incubator conditions and imaged in fresh, phenol red-free Opti-MEM as described previously .
Small interfering RNA (siRNA) transfections
Pre-validated siRNA duplexes were purchased from Life Technologies. Single-cell suspensions were prepared in proliferation media and sufficient cells seeded into 1-well of a 6-well plate for all test and control assays. Transfections were achieved using Fugene HD transfection reagent (Roche) as per the manufacturer’s protocol. Plates were returned to the incubator until cells were used for assay. Prior to assay, cells were re-suspended as single cells, counted and seeded into the NCFA as described previously. Pre-screening of knock-down efficiency found that knock-downs of both Nox2 and SOD2 were optimal (~ 50%) at 2 days using 30 μM siRNA for Nox2 and 50 μM for SOD2 (example western blot images of knock-downs are shown in Sup Fig 1). Therefore, all assays began 2 days post-treatment with these concentrations of siRNA and the equivalent of non-silencing control (Life Technologies). At the start of every assay, a dot blot was performed on cells from the starting culture to confirm the knock-down had been consistent. Quantifications of these quality control spots are shown in Sup Fig 1.
Statistical analyses were accomplished using GraphPad Prism 5 statistical software. Students t tests were used for comparison of two parameters and ANOVA or Kruskal–Wallis analyses used for > two parameters. Where significant differences were found, Dunnett, Bonferroni, or Dunn tests were used for multiple comparisons of one-way, two-way, and non-parametric ANOVA, respectively. p < 0.05 was used as the cutoff for significance and ≥ 95% statistical power. All stated “n” values indicate independent repeats.
N1 and N2 alter NSC growth
To assess the ability of the soluble N-terminal fragments to influence NSC growth, adult NSCs (harvested from wild-type mice at 8 weeks of age) were incubated with 1 μM synthetically produced N1 and N2 peptides (Fig. 1a) included in the matrix of a neural colony forming assay (NCFA; Fig. 1b). The 1 μM concentration of each peptide was based upon our previous studies showing functionality at these concentrations [33, 41, 46]. Exposure of the NSCs to either the N1 or the N2 fragment resulted in decreased growth, reducing the number of colonies formed (Fig. 1c) and, for the N2 fragment, the diameter of the colonies (Fig. 1d). The N2 growth reduction was accompanied by a significant reduction in protein expression of the cell proliferation regulatory protein Pin1 (Fig. 1e, f). Using further synthetic fragments corresponding to shorter regions of the N2 fragment, and also an N2 peptide with mutated N-terminal residues, it was found that wild-type full-length N2 is the minimum needed to cause overall growth change, especially with respect to colony formation (Fig. 1g, h). A fragment representing the most N-terminal region, amino acids 23–50, could elicit a small effect on colony diameter, but, whilst this was significantly different from the untreated control, it was not statistically different from a scrambled control peptide suggesting an artefactual result (Fig. 1h). The consequence of N-terminal mutation was less clear. Whilst the effect of wild-type N2 on reducing the number of colonies formed was clearly abolished by mutation of the N-terminal proline residues to alanine (P26/28A), there was a persisting capacity to reduce colony diameter similar to that observed with un-mutated N2 fragment.
During prion disease, ongoing PrP expression is an absolute requirement for pathogenesis . Therefore, we investigated whether the influence of the N1 and N2 peptides on cell growth was changed in cells of differing PrP expression levels. The effect of the fragments, especially for N2, did not depend upon the underlying expression of PrP in the NSCs, with both the prnp knock-out (KO) and Tga20 over-expressing cells showing reduced growth when the peptides were included in their matrix (Fig. 1i, j). However, in contrast with the wild-type cells, the KO and Tga20 cells demonstrated a changed influence of the N1 peptide, with colony diameter more influenced than the number of colonies formed.
N1 and N2 reduce migration and neurite outgrowth
Other processes that occur following division in actively replicating NSCs include migration of cells to their site of integration and the extension of neurite outgrowths, and both of these processes have been found to be influenced by cellular PrP expression levels [48, 49]. Congruent with the colony forming assay results, both migration and neurite outgrowth were reduced by the N1 and N2 peptides (Fig. 1k–m). By observing the migration of cells from the neurospheres for longer, it was apparent that the inhibitory effects of N1 and N2 were transient with migration of the N1-treated cells indistinguishable from control cells and migration resumed, albeit at an attenuated level, for N2 by 7 days (Fig. 1k).
N1 and N2 do not cause cytotoxicity or senescence
N1 and N2 reduce intracellular ROS production
To directly investigate the role of Nox signalling, siRNA knock-down was used to target Nox2. Nox2 was selected as it is a central catalytic subunit of the Nox complex and known to have a role in the maintenance of NSC populations within the brain . Nox2 has also previously been linked with PrP signal transduction and to prion disease pathogenesis [51, 52, 53]. Western blotting confirmed that the stem cells do express this subunit and showed no significant changes in the expression levels of Nox2 after 24 h incubation with N1, N2 or DPI (Fig. 3f, g). Confocal imaging of PrP and Nox2 showed altered cellular morphology and localisation of Nox in response to the peptides (Fig. 3h). Using siRNA to knock down the expression of Nox2, a significant decrease in cellular growth was seen in the NCFA (consistent with the previously reported effects of DPI treatment) and this completely negated the effects of both N1 and N2 (Fig. 3i, j).
Reduced redox signalling causes increased mitochondrial fission
N1 and N2 cause changes in SOD2 expression and SOD activity
NSCs show altered PrP N-terminal detection depending upon whether they are actively growing or quiescent
Difficulty assigning a single or predominant function to PrP may have arisen, at least in part, from the numerous post-translational modifications to which it is subjected and the biochemically distinct protein/peptide species that are produced as a result. The data presented herein show that the N-terminal cleavage fragments, N1 and N2, can reduce intracellular redox signalling, with the N2 peptide directly reducing Nox catalytic activity. The reduced ROS increases the activity of DRP1, which mobilises to the mitochondria resulting in their fission within a few hours and later increasing protein levels of SOD2. The increased SOD2 levels and activity in turn maintain a low ROS environment within the mitochondria preserving cells in a quiescent state (Fig. 8).
The enzymes responsible for N1/C1 cleavage are still a matter of contention. It was first reported that A Disintegrin and metalloproteinase domain-containing protein 10 (ADAM10) was the candidate alpha-cleavage enzyme , although later work found that this enzyme was primarily responsible for secretory cleavage of PrP near the GPI anchor . ADAM17 (alternatively known as tumour necrosis factor-α-converting enzyme [TACE]) and ADAM9 have also been implicated in N1/C1 cleavage, but a definitive identification has not yet been made; a detailed review of this work can be found in . The N2/C2 cleavage event has been demonstrated to be mediated by ROS  and, during scrapie infection, by the calpain family . Conditions that change the expression or activity of cleavage enzymes may shift the balance between full-length PrP and its cleavage fragments or shift the ratio of alpha- and beta-cleavage resulting in changed cellular outcomes. Therefore, the apparent cellular function of PrP may be as related to cellular protease expression levels as it is to total PrP levels.
Alternative explanations may exist for the observed differences in biological effects between the N1 and N2 fragments. The N1 fragment contains a second charged cluster domain (resulting in charged domains at both ends of the peptide); this may endow it with extra properties such as the ability to bind and engage pathways that N2 cannot. Such putative-binding capacity may result in less N1 being available for modulation of the tested pathways. We have also observed aggregation of the 23-89P26/28A mutant in the previous studies  and the second charged domain within N1 may facilitate greater dimerisation/oligomerisation as compared with N2 thereby rendering less N1 peptide effectively available for interactions. Furthermore, only a single peptide concentration was compared in the current study; therefore, it is possible that higher concentrations of N1 could elicit the same intensity of reaction of the tested concentrations of N2. It is also unclear from the presented data why a difference should exist in cellular responses to N1 as a result of endogenous PrP expression; this could be related to the basal levels of N1 within these cells or lack of N1 and/or the pathways compensating for changed expression levels of the full-length protein. As the knock-out and Tga20 over-expressing NSCs demonstrate changed growth properties from the wild-type, which is especially apparent for the number of colonies generated by the Tga20 cells, basally changed pathways could influence how N1 is able to interact with and modulate cellular growth pathways.
PrP has been linked with increased ROS generated by Nox in the context of prion disease  and oxidative damage caused by overstimulation of Nox signalling was found to be mediated by interaction of the N-terminus at the cell membrane in an antibody-stimulated model of toxicity . Tethering the N-terminus of PrP to the cell membrane reduces the capacity of PrP to undergo normal N2/C2 cleavage, as well as cellular resistance to oxidative stress , possibly by preventing the N2 fragment from modulating Nox function. How exactly the N-terminal fragments or N-terminus of full-length PrP interact with Nox is unknown. Both our previous in vitro studies that indicated no intrinsic antioxidant activity of N2  and, herein, the decreased utilisation of NADPH substrate in response to N2 argue against an indirect ‘mopping up’ of the superoxide radicals. Other possibilities include direct interaction with one or more of the Nox subunits or modulation of the lipid membrane environment in which the Nox complex forms. The latter is a reasonable possibility as we have shown that both of these peptides bind to lipids under specific conditions [64, 65] and changes in the lipid environment are observed during N2 protection against oxidative stress .
From the data presented, we cannot rule out a direct effect of the peptides on the mitochondria themselves. Whilst N1 did not significantly change Nox activity or cellular SOD2 expression it did alter mitochondrial fission and knocking down SOD2 completely counteracted the effect of this peptide on cellular growth. Of interest, it has recently been reported that a population of PrP resides in the mitochondria of normal disease-free cells and is endoproteolytically cleaved similar to total brain PrP . This might offer the N-terminal peptides, especially N1 as the predominant N-terminal cleavage product, unhindered access to the mitochondria from inside the cell, directly facilitating an influence of the peptide on this organelle. Consequently, this might also have a bearing on the apparent lower efficacy of N1 in stimulating these responses. We have previously shown that N2 traffics from the cell surface to the mitochondria , but this has not, to our knowledge, been examined for N1. The N1 fragment may be more functionally effective in stimulating quiescence if generated at the site of action (the mitochondria) rather than when it requires trafficking from the cell surface.
The role of SOD2 in regulating the cell cycle has become well characterised [67, 68]. Increased cellular levels of SOD2 are known to facilitate cellular transition to quiescence, whereas loss of SOD2 protein or increased superoxide signalling favours increased cellular growth [69, 70]. PrP has additionally been linked with SOD2 in the context of prion pathogenesis. During prion disease or prion infection of cells, changes of expression and activity of the SOD family of enzymes are seen, including a decrease in SOD2 protein levels [45, 71]. Correspondingly, neurogenesis during prion disease increases . A further point to note when considering an influence of PrP-mediated signalling on SOD2 is that aberrations in SOD2 expression are linked with the uncontrolled cell cycling of cancer cells [68, 72]. Recently PrP expression has been linked with several cancers as well as the risk of metastases [73, 74, 75]. In addition to changes in mitochondrial SOD2, changes in mitochondrial fission and fusion have been linked with the uncontrolled cell cycling during cellular adaptation to the energy demands of cancer . Regulation of SOD2 expression and mitochondrial dynamics by the N-terminal cleavage fragments of PrP or putatively the N-terminus in association with the full-length protein may explain why such disparate maladies are linked with this protein.
From our data, it would appear that detection of full-length PrP is predominantly observed in proliferating cells or cells residing alongside those that are actively proliferating, with quiescent cells staining only for C-terminal PrP. This might suggest that the signals stimulated by the N-terminal fragments can function in an autocrine or paracrine manner. Alpha-cleavage of PrP has been demonstrated to increase when PrP homodimerises at the cell membrane  and this may represent a mechanism for initiating intra/intercellular signalling. Questions remain as to the actions of full-length PrP, secretory (GPI-anchorless) PrP, C1, C2, or gamma cleaved PrP fragments in this pathway or cross-interactions with N1 and N2, but it seems highly likely that these proteins will be functional in their own right, bringing many more layers of complexity to the PrP function narrative.
Cross-talk between cellular redox signalling pathways and the mitochondria, and between the mitochondria and the nucleus, have been the subject of much study, but the role of the PrP N-terminal cleavage fragments as upstream modulators of these pathways has not previously been considered. As increased PrP expression levels are linked with increased growth of NSCs and herein we have linked the N-terminal cleavage fragments with reduced cell growth/increased cell quiescence, dynamic modulation of PrP cleavage may be part of an important cellular homeostatic mechanism. The modulation of redox signal transduction appears to be the first event influenced by the PrP cleavage fragments, after which the reduced intracellular ROS indicates that cells should enter a resting phase of life as modulated and maintained by their mitochondria. The diversity of the potential outcomes resulting from up-steam regulation of central redox signalling pathways in different cell types could have wide-reaching implications for many of the reported functions of PrP as this small protein begins to reveal its functional complexity.
SJC is funded by an NH&MRC Practitioner Fellowship (#APP1105784). SCD is supported by a senior research fellowship administered by the faculty of Medicine, Dentistry and Health Sciences, University of Melbourne, and CLH was the recipient of a CJDSGN Rhonda McCoy memorial fellowship. This research was supported in part by the Intramural Research Program of the NIH, [NIAID].
Compliance with ethical standards
Conflict of interest
The authors declare that they have no conflicts of interest.
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