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Astrocytes migrate from the optic nerve into the inner retina, forming a template upon which retinal vessels develop. In the Nuc1 rat, mutation in the gene encoding βA3/A1-crystallin disrupts both Notch signaling in astrocytes and formation of the astrocyte template. Here we show that loss of βA3/A1-crystallin in astrocytes does not impede Notch ligand binding or extracellular cleavages. However, it affects V-ATPase activity, thereby compromising acidification of the endolysosomal compartments, leading to reduced γ-secretase-mediated processing and release of the Notch intracellular domain (NICD). Lysosomal-mediated degradation of Notch is also impaired. These defects decrease the level of NICD in the nucleus, inhibiting expression of Notch target genes. Overexpression of βA3/A1-crystallin in those same astrocytes restored V-ATPase activity and normal endolysosomal acidification, thereby increasing the levels of γ-secretase to facilitate optimal Notch signaling. We postulate that βA3/A1-crystallin is essential for normal endolysosomal acidification, and thereby, normal activation of Notch signaling in astrocytes.
Notch signaling is involved in numerous functions including pattern formation during development and gliogenesis 1, 2. The Notch pathway is evolutionarily conserved and promotes the generation of astrocytes from neuronal precursor cells 3, 4. Four Notch receptors, Notch1-4, have been identified. The Notch gene encodes a single-pass transmembrane protein which appears on the cell surface as a heterodimeric protein after maturing along the secretory pathway 5. Notch receptors are activated by interaction with transmembrane ligands expressed on the surface of juxtaposed cells. Binding of the ligands activates two sets of proteolytic cleavages in the Notch receptor. First, TNFα-converting enzyme (TACE), a member of the ADAM family of matrix metalloproteases, cleaves the receptor at its extracellular domain, generating a membrane-tethered Notch receptor 5. Next, the receptor is cleaved within its transmembrane domain by the γ-secretase multiprotein complex (including presenilin, nicastrin, APH1 and PEN2) 6, resulting in release of the Notch intracellular domain (NICD); NICD translocates to the nucleus to activate Notch target genes 7.
The activation of Notch requires processing, endocytosis and trafficking of both Notch receptor and ligand. Ubiquitination has an important role in the activation of Notch signaling 8. In recent years, it has also been suggested that endocytosis of Notch is important for ligand-mediated activation 9, 10. There is now a general consensus that γ-secretase-mediated S3 cleavage of Notch can occur at multiple steps within the endolysosomal compartments 11. Increasing evidence suggests that endocytosis is involved in the transport of ligand-receptor complexes to acidic endolysosomal compartments, where low pH is required for signaling 10, 12. The acidic luminal environment of endosomes, lysosomes and secretory vesicles is established by the vacuolar-type proton ATPase (V-ATPase), which pumps protons into the lumen 13–15. V-ATPase-dependent acidification is required to optimally activate γ-secretase, an acid protease that mediates S3 cleavage of Notch 15. Therefore, progressive acidification of the endolysosomal system could influence the levels of NICD production. In Drosophila, various mutants that show impaired acidification due to alteration of V-ATPase activity affect Notch signaling 13, 15.
In the past decade, several laboratories have shown that crystallins, the major structural proteins of the lens, are expressed outside of the lens and have cellular functions 16–18. We previously postulated that βA3/A1-crystallin, may regulate the process of programmed cell death in the developing retina and that it also plays a pivotal role in maintaining the proper number of astrocytes in the retina through an anoikis-mediated cell death process 19, 20. We show here that βA3/A1-crystallin affects Notch signaling in astrocytes by regulating endolysosomal function in astrocytes by modulating the activity of V-ATPase. In the absence of βA3/A1-crystallin, V-APTase activity decreases and acidification of endolysosomal compartments is impaired. The higher pH inhibits the γ-secretase-mediated formation of NICD, leading to decreased Notch signaling.
Notch signaling plays an important role during differentiation and maturation of brain astrocytes 21, 22; however, its role in optic nerve astrocytes is unexplored. Disrupted Notch signaling affects pattern formation in other systems 23. We have shown major defects in template formation and honeycomb patterning by Nuc1 retinal astrocytes with mutated βA3/A1-crystallin 24, 25. We first studied expression of Notch pathway components in optic nerve astrocytes cultured from 2–3 day old WT (wild type) and Nuc1 rats. Quantitative reverse transcriptase PCR (qRT-PCR) analysis showed that WT cells strongly express Notch1 receptor transcript (Figure 1a) with trace levels of Notch4 (data not shown). Nuc1 cells have 2 fold higher expression of Notch1 than WT cells. The canonical Notch ligands are single pass transmembrane proteins belonging to the Delta-like or serrate families. Using specific primers, we tested their expression in WT and Nuc1 astrocytes. Our results show, for the first time, that optic nerve astrocytes express Jagged1 (Figure 1b, c), with no significant expression of the Delta-like ligands. Co-localization of Jagged1 with the astrocyte marker, glial fibrillary acidic protein (GFAP), in retinal flatmounts confirms expression of Jagged1 in retinal astrocytes (Figure 1d).
Next, we investigated the expression of NICD using anti-Notch V1744, an antibody specific for NICD. Immunostaining and western blotting detected a significant decrease in NICD protein in Nuc1 astrocytes compared to WT (Figure 1e, f). Confocal microscopy (Figure 1g) demonstrated that NICD (red) staining was primarily nuclear and weaker in Nuc1 astrocytes compared to WT, while Jagged1 (green) staining was similar in both cell types.
Since our data indicate downregulation of NICD in Nuc1 astrocytes, we examined activation of the Notch pathway using a Notch-luciferase reporter assay. Notch1 receptor constructs, including a full-length receptor (Notch-FL) and truncated or mutant derivatives with varying activation profiles, were used. A constitutively active Notch construct, NδE, activates the Notch pathway in a ligand-independent manner, and the NICD construct contains only the Notch intracellular domain. WT and Nuc1 astrocytes were co-transfected with Notch-FL, NδE or NICD constructs, along with the 6XCBF1-luciferase Notch reporter plasmid, and the phRL-SV40 Renilla luciferase reporter plasmid. WT astrocytes, transfected with the NotchFL construct, showed 4-fold increased activity compared to astrocytes transfected with vector only (Figure 2a). Nuc1 astrocytes, in contrast, showed only minimal activation (Figure 2b). The Notch pathway was activated equally in WT and Nuc1 astrocytes expressing the constitutively active NδE construct (Figure 2a, b). Furthermore, similar activation was observed in WT and Nuc1 astrocytes expressing the NICD construct (not shown). Notch1 in WT astrocytes was strongly activated (~65% increase in luciferase activity compared to untreated cells) by EDTA, a Ca2+-chelating agent which activates Notch in a ligand-independent manner 26. This activation was inhibited by N-[N-(3, 5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT), an inhibitor of γ-secretase (Figure 2a). Nuc1 astrocytes, however, showed only modest activation of the Notch pathway by EDTA, although DAPT did inhibit this Notch activation (Figure 2b). EDTA did not significantly influence Notch activity in either WT or Nuc1 astrocytes expressing NδE, although DAPT still was inhibitory (Figure 2a, b). DMSO (vehicle for DAPT) did not affect Notch activity in WT or Nuc1 astrocytes.
We also assessed the expression of Notch target genes using qRT-PCR. Expression of Hes1 and Hey1, was reduced ~65% and ~75% respectively; in Nuc1 astrocytes compared to WT astrocytes (Figure 2c, d).
Our studies have shown that optic nerve astrocytes strongly express Jagged1. Previous studies have shown that Jagged1 acts as an inhibitor of the Notch signaling pathway by competitively inhibiting binding of Delta-like ligands to the Notch receptor 27, 28. Therefore, we sought to determine whether Jagged1 acts as agonist or antagonist in astrocytes. WT and Nuc1 astrocytes were treated with Jagged1 siRNA, and Notch pathway activity monitored by measuring expression of Hey1 and Hes1. siRNA knockdown of Jagged1 (Figure 3a) resulted in concomitant reductions in Hes1 and Hey1, in both WT and Nuc1 astrocytes (Figure 3b).
To study the Jagged1-mediated activation of Notch signaling, we used a co-culture system, where cells expressing full length Notch1 receptor and the 12XCSL-luciferase Notch reporter and pRL-CMV renilla plasmids were interfaced with cells expressing full length Jagged1. We observed robust 12XCSL-luciferase activation in WT cells, whereas Nuc1 cells showed much weaker activation of the Notch pathway (Figure 3c). As control, a vector expressing a mutant Jagged1 protein that fails to bind to the Notch receptor (Jagged1ΔEGF) was used. This resulted in an abrogation of Notch signaling, suggesting that the trans-interactions with Jagged1 expressing cells were inhibited.
Since optic nerve astrocytes express both Notch1 and Jagged1, we next asked whether the Notch receptor is cis-inhibited by ligand expressed in the same cell and trans-activated by ligand expressed on neighboring cells. In most cancer cells, Notch receptor and ligand are expressed by the same cell, with the ligand inhibiting the receptor, thereby facilitating the receptor trans-interaction with ligand expressed by neighboring cells 29. We studied the trans-interactions by silencing Jagged1 expression using siRNA and transfecting the same cells with the 12XCSL-luciferase Notch reporter plasmid and pRL-CMV Renilla plasmid. Figure 3d shows significant induction of Notch signaling in both WT and Nuc1 astrocytes, suggesting a cis-inhibition of the receptor by Jagged1 in the same cell and trans-activation by Jagged1 on neighboring cells. Cells transfected with non-targeting siRNA were also co-cultured with Jagged1 overexpressing cells, showing only a modest induction of the Notch pathway in WT, and none in Nuc1 astrocytes (Figure 3d).
Our data show that NICD is decreased in Nuc1 cells despite the higher levels of Notch1 receptor. Since this could result from defective proteolytic cleavage of Notch, we analyzed the two cleavage steps required for Notch activation. The S2 and S3 cleavages are mediated by ADAM metalloprotease and γ-secretase, respectively 5. In cells treated with a hydroxamate ADAM inhibitor, BB-94, the mRNA expression of Notch targets Hes1 and Hey1 was inhibited in both WT and Nuc1 cells (Figure 4a). To confirm cleavage of the Notch receptor, we used a myc-tagged, constitutively active, membrane-bound derivative of Notch1 (NδE). Cleavage of NδE at V1744 by γ-secretase generates a product corresponding to NICD. This construct was visualized by immunoblotting with anti-Myc antibody. In both WT and Nuc1 optic nerve astrocytes expressing NδE, an approximately 85-kDa protein representing uncleaved NδE and a 75 kDa band corresponding to NICD were detected.
To confirm the identity of these fragments, WT and Nuc1 astrocytes were first transfected with a NδE construct mutated at residue 1744 (V1744K) to prevent cleavage and generate a band co-migrating with the higher-molecular-mass NδE band. Further, astrocytes transfected with an NδE-derived construct truncated at the cleavage site (ICv1744), produced a fragment co-migrating with NICD (Figure 4b&c). Since both the cleaved and uncleaved forms were generated in Nuc1 astrocytes expressing NδE, the cleavage sites of Notch1 are functional in the mutant astrocytes.
We next investigated whether nuclear translocation of NICD is inhibited in Nuc1 astrocytes. For this, we overexpressed NICD in WT and Nuc1 astrocytes using a constitutively active Notch1 construct in a pCDNA3.1 plasmid. Overexpression was confirmed by western analysis (Figure 5a). Increased staining for NICD was seen in both WT and Nuc1 astrocytes compared to astrocytes transfected with vector alone (Figure 5b). qRT- PCR showed that overexpression of NICD by Nuc1 astrocytes strongly rescued expression of Notch target genes (Figure 5c). The expression of Hey1 and Hes1 was also measured in WT and Nuc1 cells transfected with Notch-FL and Notch extracellular truncation (NEXT) plasmids. In contrast to overexpression of NICD, neither of these constructs rescued Hey1 or Hes1 expression in Nuc1 astrocytes (Figure 5d, e).
γ-Secretase is enzymatically more active in the low pH environment of the endolysosomal compartments 9, 30. In the case of Amyloid Precursor Protein, a well characterized γ-secretase substrate, cleavage to produce β amyloid occurs in the endolysosomal compartments 31. To determine if the activity and sub-cellular distribution of γ-secretase is aberrant in Nuc1 astrocytes, post-nuclear supernatants from WT and Nuc1 astrocytes were layered on iodixanol gradients and subjected to ultracentrifugation. The composition of the fractions was determined by immunoblotting with specific markers. Lysosome-associated membrane protein1 (LAMP1)-positive lysosomes were strongest in fractions 3, 4, and 5; Rab5-positive endosomes in fractions 6, 7, and 8; and Golgi in fractions 9 and 10 (Figure 6a). The activity of γ-secretase was measured by performing ELISA to quantify production of β-amyloid 40. In WT astrocytes, γ-secretase activity (Figure 6b) was present in fractions 3, 4, and 5 corresponding to the lysosomes; fractions 6, 7, and 8 corresponding to the Rab5-positive endosomes. We also detected modest activity in the Golgi. Nuc1 astrocytes showed significantly decreased γ-secretase activity in the endolysosomal compartments, whereas activity in the Golgi was relatively unchanged (Figure 6c). Overexpression of βA3/A1-crystallin in Nuc1 astrocytes restored normal γ-secretase activity (Figure 6d). To confirm that βA3/A1-crystallin modulates γ-secretase activity in astrocytes, we performed Cryba1 siRNA silencing studies to downregulate βA3/A1-crystallin in WT rat astrocytes in culture using specific ON-TARGET plus SMART pool siRNA (Dharmacon, CO). Scrambled siRNA was used as control. βA3/A1-crystallin downregulation was confirmed at the mRNA level by quantitative PCR analysis and at the protein level by immunoblotting (data not shown). Knockdown of βA3/A1-crystallin in WT astrocytes decreases γ-secretase activity, but the decrease is not as robust as in Nuc1 homozygous astrocytes, the activity could be rescued to normal levels by subsequent overexpression of βA3/A1-crystallin (Supplementary figure S1). As proof of principle that βA3/A1-crystallin can modulate γ-secretase activity in astrocytes, we used Nuc1 heterozygote astrocytes in culture and downregulated βA3/A1-crystallin with siRNA as above. The concentration of siRNA required to downregulate Cryba1 in Nuc1 heterozygotes was significantly lower than that needed with WT astrocytes. Heterozygote astrocytes in which Cryba1 was downregulated showed loss of γ-secretase activity to levels similar to that in Nuc1 homozygous astrocytes. Activity was rescued by overexpression of βA3/A1-crystallin in the same cells (Figure 6e). In addition, Cryba1 floxed mice were generated (Supplementary figure S2); downregulation of βA3/A1-crystallin in astrocytes cultured from these mice, using an adenoviral vector expressing Cre-recombinase, also decreased γ-secretase activity, however, this activity was rescued by subsequent over-expression of βA3/A1-crystallin (Figure 6f). γ-Secretase activity was also decreased in endolysosomes of both WT and Nuc1 astrocytes pre-treated with chloroquine, an inhibitor of lysosomal acidification (Supplementary figure S3). The decrease in activity was more significant in WT astrocytes than in Nuc1. γ-Secretase activity was not affected in Golgi, a non-acidic compartment. Previous studies, and our results, indicate a stringent pH requirement for γ-secretase activity 12, 31, 32. While low levels of γ-secretase activity are detectable in Nuc1 astrocytes, that activity may be insufficient for optimal Notch signaling. Previous studies in other systems have shown that treatment with DAPT resulted in a marked decrease in the generation of NICD, as monitored by expression of Hey1 and Hes1 5, 33. qRT-PCR analysis revealed that DAPT treatment reduced Hes1 and Hey1 similarly in WT and Nuc1 cells (Supplementary figure S4).
Since γ-secretase activity is compromised in astrocytes lacking functional βA3/A1-crystallin, we suspected that endolysosomal acidification might be dysregulated. To obtain direct evidence that this may be the case, we compared V-ATPase activity in astrocytes lacking βA3/A1-crystallin with WT. Isolated lysosomes were incubated in acridine-orange and accumulation of protonated acridine in the lysosomes measured by the resultant fluorescence. In the presence of exogenous ATP, H+ uptake was significantly increased in the lysosomes of WT astrocytes. Under the same conditions, the acridine orange fluorescence in Nuc1 astrocytes was significantly lower (Figure 7a). V-ATPase activity in Nuc1 homozygote astrocytes was rescued by βA3/A1-crystallin overexpression (Figure 7b). We further demonstrate reduced V-ATPase activity in βA3/A1-crystallin-downregulated Nuc1 heterozygote astrocytes (Figure 7c), and in astrocytes isolated from Cryba1 floxed mice, after βA3/A1-crystallin deletion (Figure 7d). Moreover, the V-ATPase activity could be rescued by subsequent overexpression of βA3/A1-crystallin (Figures 7c and 7d). As control, cells were treated with the V-ATPase inhibitor (bafilomycin a1) and significant inhibition of lysosomal V-ATPase activity was observed (data not shown).
Endolysosomal pH was measured by monitoring fluorescence changes in astrocytes loaded with FITC-dextran. In WT the average endolysosomal pH was 4.5, increasing to 5.7 in Nuc1 astrocytes (Figure 7e). This increase in pH was reversed by overexpression of βA3/A1-crystallin in the same cells (Figure 7f). Similarly, loss of βA3/A1-crystallin in Nuc1 heterozygote and Cryba1-floxed astrocytes induced alkalinization by ~1pH unit and was rescued to near normal levels by subsequent overexpression of βA3/A1-crystallin in the same astrocytes (Figure 7g&h), suggesting a direct role of βA3/A1-crystallin in regulating acidification of astrocytes.
Since defective acidification of endolysosomal compartments could also lead to impaired degradation of Notch, we compared the clearance of the Notch receptor in WT and Nuc1 astrocytes after transfection with the Myc-tagged full length Notch1 receptor. To activate the Notch pathway, these cells were co-cultured with cells overexpressing Jagged1. After 1 hour, the myc-tagged Notch1 underwent significant clearance from the cytosol in WT astrocytes. In Nuc1 astrocytes, however, Notch1 strongly persisted. After 3 hours, most of the tagged receptor was degraded in WT, but much remained in the perinuclear region of Nuc1 astrocytes (Figure 8a).
Since degradation of Notch1 appeared dysfunctional in Nuc1 astrocytes, we performed subcellular fractionation using sucrose density gradients. Notch1 was found at interfaces 1 and 2, containing LAMP1 positive lysosomes and Rab5-positive early endosomes, respectively (Figure 8b). The Nuc1 homozygote astrocytes showed greater accumulation of Notch1 in these compartments than did WT astrocytes, suggesting that Nuc1 cells have a compromised ability to degrade and process the Notch receptor. Moreover, overexpression of βA3/A1-crystallin in Nuc1 homozygote astrocytes restored normal lysosomal-mediated degradation of Notch (Figure 8b).
Since Nuc1 results from a mutation in βA3/A1-crystallin, we also determined the subcellular localization of βA3/A1-crystallin in astrocytes. Immunostaining revealed that while βA3/A1-crystallin was predominantly cytosolic in both WT and Nuc1 astrocytes, it also co-localized with LAMP1-positive lysosomes in WT astrocytes and, to a lesser extent, in Nuc1 astrocytes (Figure 8c). Western analysis of the sucrose gradient fractions confirmed these results (Figure 8d).
Our results demonstrate that βA3/A1-crystallin is required for normal lysosomal acidification. To determine if βA3/A1-crystallin also modulates the activity of lysosomal enzymes, we assayed the activity of Cathepsin D, a lysosomal hydrolase requiring low pH for optimal activity. The intermediate chain of Cathepsin D is processed into the active heavy and light chains once the protein encounters the acidic lysosomal environment. Changes in this processing reflect altered function of lysosomal hydrolases. Levels of both the heavy and light chains were decreased in Nuc1 homozygote astrocytes (Figure 8e) and Cathepsin D activity was rescued by overexpression of βA3/A1-crystallin (Figure 8f). Our data also indicate that in Nuc1 heterozygotes and in Cryba1-floxed astrocytes, Cathepsin D activity was markedly reduced, while subsequent overexpression of βA3/A1-crystallin rescued activity (Figure 8g&h).
Functional effects of Cryba1 knockout from Cryba1-floxed mouse astrocytes are demonstrated above (Figures 6–8). To prove that Notch signaling is affected, we measured Hes1 and Hey1 expression, finding it decreased in the Cre-infected astrocytes and rescued by subsequent overexpression of βA3/A1-crystallin (Supplementary figure S5).
To determine whether compromise of the ubiquitin-proteosome system in Nuc1 astrocytes might affect Notch degradation, we measured the intracellular levels of the receptor in the presence of specific inhibitors of the ubiquitin proteosomal pathway, clasto-lactacystin β-lactone and MG132 34. Treatment with neither inhibitor resulted in a significant accumulation of Notch1 receptor in WT or Nuc1 homozygote astrocytes (Supplementary figure S6). Thus, our results suggest that in astrocytes, endogenous Notch receptor is posttranslationally regulated predominantly by the lysosomal-mediated degradative pathway.
Notch signaling has been implicated in the demethylation and activation of the GFAP promoter 35. Since GFAP is an important player in astroglial differentiation 35 and since the astrocyte template in the Nuc1 homozygote retina is abnormal, we wondered if Nuc1 astrocytes differentiated normally. To determine if Notch is involved in the transcriptional regulation of GFAP in optic nerve astrocytes, we assayed GFAP promoter activity using a GFAP promoter-reporter construct containing the rat GFAP promoter upstream of the firefly luciferase cDNA and the Renilla luciferase reporter under the control of CMV promoter construct. WT and Nuc1 astrocytes were transfected with these constructs and luciferase activity measured as readout for GFAP promoter activity (Supplementary figure S7). WT astrocytes showed a much stronger GFAP promoter activity than did Nuc1 astrocytes. To test whether GFAP promoter activation is due to Notch signaling, we treated cells with DAPT prior to transfecting the cells with the promoter-reporter constructs. Both WT and Nuc1 astrocytes showed a reduction in GFAP reporter activity, confirming the role of the Notch pathway in differentiation of optic nerve astrocytes.
The retina requires two macroglial cells, Muller cells and astrocytes, for proper functioning. While Muller cells are retina-specific, astrocytes are present throughout the CNS, and migrate from the optic nerve head into the inner retina 36. Our studies with the Nuc1 spontaneous mutant rat, and the work of others, have demonstrated that, in rodents, an astrocyte template is essential for the proper formation of the retinal vasculature 24, 37–39. No signaling mechanisms have yet been shown to regulate formation of this template. Notch is the most significantly enriched signaling pathway in developing tissues and is known to regulate pattern formation in other systems; its dysfunction results in various developmental defects and adult pathologies 40.
We show that optic nerve astrocytes express both the Notch receptor and its ligand, Jagged1. This suggests that in astrocytes, Notch is involved in juxtacrine signaling, where Jagged1 from one astrocyte activates the Notch receptor on a neighboring astrocyte. This juxtacrine signaling may be critical to the proper organization of astrocytes in the developing retina. In Nuc1, defective Notch signaling may account for improper template formation and patterning by retinal astrocytes.
Notch signaling is known to play a pivotal role in cell fate specification during development. For example, it has been shown that blocking Notch signaling causes fewer Muller glial cells to be made in the retina, while an expansion of Muller glial cells was observed when Notch was activated 41, 42. Decreased Notch signaling in Nuc1 astrocytes may account for the abnormal morphology and decreased GFAP, that we have reported 24, 43. We now show lower gene promoter activity for GFAP in Nuc1 astrocytes, which might potentiate the decreased astrocyte number and abnormal template formation seen in the retinas of Nuc1 rats 24, and in transgenic mice overexpressing the mutant βA3/A1-crystallin protein 43.
We have previously shown that in retinal pigmented epithelial cells, βA3/A1-crystallin is trafficked to the lysosomes and plays a pivotal role in the degradation of phagocytized photoreceptor outer segments 44. This prompted us to investigate whether βA3/A1-crystallin modulates Notch signaling by manipulating lysosomal pH in astrocytes. Luminal acidification of endolysosomal compartments is regulated by a major proton pump, V-ATPase 45. We show that βA3/A1-crystallin may affect endolysosomal acidification by regulating the activity of V-ATPase. Although organelles use the same V-ATPase to acidify their interiors, each is able to maintain a stable, characteristic internal pH 46. V-ATPase activity is necessary for proper activation of Notch signaling in the endosomal system, although it is not required for Notch to access endosomal compartments 13. In astrocytes, in which βA3/A1-crystallin is non-functional, the activity of V-ATPase is compromised, diminishing acidification of endolysosomal compartments and resulting in decreased Notch activation. This is consistent with studies showing that normal functioning of V-ATPase is required for optimal catalytic activity of γ-secretase 45, 47. Loss of functional βA3/A1-crystallin in astrocytes does not impede Notch ligand binding nor extracellular cleavages, because V-ATPase activity is only required after ligand binding. However, loss of βA3/A1-crystallin affects the γ-secretase-mediated processing of NICD in astrocytes. The resultant dysfunction, affecting release of NICD, inhibits transcription of Notch target genes.
Lysosomes are cellular organelles primarily involved in degradation and recycling processes 48. V-ATPase also plays an important role in the lysosomal degradation of Notch 14. Again, the regulation of proper luminal pH is critical for this process, since the lysosomal hydrolases that degrade internalized proteins, lipids and complex carbohydrates function optimally only in the acidified lysosomal microenvironment 15. Our data on Cathepsin D, one such lysosomal hydrolase, is consistent with a general decrease in lysosomal hydrolase activity in astrocytes lacking βA3/A1-crystallin. In Nuc1 astrocytes, the Notch receptor does not undergo lysosome-mediated degradation at a normal rate, resulting in its abnormal accumulation. Similar results are seen in cells exposed to chloroquine, an inhibitor of lysosomal acidification 34. Since ligand-independent Notch activation does not normally occur in mammalian cells when Notch degradation is impaired 49, acidification of the endolysosomal compartment in astrocytes is essential both for Notch signaling and degradation, suggesting a novel role for βA3/A1-crystallin as an important regulator of this acidification process.
In conclusion, our study describes an unanticipated function for βA3/A1-crystallin in endolysosomal acidification in astrocytes as depicted in the hypothetical model (Figure 8i). Regardless of the specific mechanism whereby mutation of βA3/A1-crystallin affects the activity of V-ATPase, and thus reduces endolysosomal acidification, our data suggest that through its action in maintaining normal lysosomal pH, βA3/A1-crystallin has a fundamental effect on Notch signaling in astrocytes.
The following antibodies were used: Notch1 (SAB3300078, Sigma, St. Louis, MO; ab27526, Abcam, Cambridge, MA; H731, Santa Cruz Biotechnology Inc, Santa Cruz, CA), Jagged1 (SC-8303, Santa Cruz Biotechnology Inc), GFAP (Z0334, DAKO Corporation, Carpentaria, CA), Myc (2272S, Cell Signaling Technology, Danvers, MA), LAMP1 (H4A3-C, Developmental Studies Hybridoma Bank, University of Iowa, Iowa), Rab5 (C8B1, Cell Signaling Technology), Calnexin (ab75801, Abcam), Cathepsin D (SC-6487, Santa Cruz Biotechnology).
βA3/A1-crystallin (Cryba1) floxed homozygous mice were generated. A BAC clone (Sp6 library, SV129 origin) encompassing the mouse βA3/A1-crystallin locus was obtained through the Invitrogen BAC screening service (Invitrogen, CA) and was used to construct a conditional targeting vector in which exons 4 and 5 were flanked by LoxP sites by recombineering as described 50. The targeting vector contained a PGK driven DTA (modified diphtheria toxin A) cassette 51 as the negative selection marker, and was electroporated into W4 (129S6/SVEvTac) ES cells 52. G418-resistant clones were picked, expanded, and screened for homologous recombination by both long-range genomic PCR and Southern blotting. Construction of the targeting vector containing the cKO allele was made as depicted in Supplementary Figure 2. Correctly targeted clones were further analyzed by karyotyping. Embryonic stem cell clones carrying the targeted allele were isolated, and three clones were injected into blastocysts. The selectable marker (frt - NeoR - frt) was removed by mating to mice (Jackson Laboratories) expressing flp recombinase hACTB/FLPe (strain name: B6.Cg-Tg (ACTFLPe) 9205Dym/J) during early development. Successful excision of the selectable marker cassette was confirmed by PCR spanning the insertion site.
Monolayer astrocytes from Cryba1 floxed mice were infected with control adenovirus (Ad CMV eGFP) or Cre-recombinase virus (Ad CMV Cre-RSV GFP; KeraFAST Inc, Boston, MA). 104 cells were infected in triplicate at MOI (multiplicity of infection) 0.01–10,000 pfu/cell. Cell survival was assessed 48 hours post infection by MTT assay. Cells infected for 48 hours were transfected with pcDNA3.1-Cryba1 vector to overexpress βA3/A1-crystallin and harvested 36 hours later.
Optic nerve astrocytes from P2 rats (WT and Nuc1) and from P2 Cryba1 floxed mice were cultured in DMEM-F12 medium containing 10% FBS, as recently described 53.
2 μg of total RNA was reverse-transcribed to cDNA in a 20 μl reaction volume using superscript reverse transcription kit (Invitrogen, Carlsbad, CA). PCR amplification was performed utilizing the 7500 PCR Fast Real-Time System (Applied Biosystems, Carlsbad, CA) with TaqMan probes for Notch1 (Rn01758633_m1), Notch2 (Rn01534371_m1), Jagged1 (Rn00569647_m1), Hey1 (Rn00468865_m1), Hes1 (Rn00577566_m1), NRP1 (Rn00686106_m1), HPRT(Rn01527840_m1) and GAPDH (Rn01775763_g1*). The reaction consisted of the following steps, enzyme activation at 95°C for 20 seconds, 40 cycles of denaturation at 95°C for 3 seconds combined with annealing/extension at 60°C for 30 seconds. All data were analyzed with the ABI 7500 real time PCR system using the data assist software (Applied Biosystems); graphs were plotted using Microsoft Excel. All experiments were performed at least three times in triplicate. Data are represented as mean ± standard deviation (SD).
Western blots were done as previously described and quantified by densitometric analysis using ImageJ 54.
For Notch luciferase assays, lipofectamine 2000 (Invitrogen) was used to transfect cells with the different Notch constructs and pGL2–6XCBF-1 Notch luciferase reporter construct (2.3 μg) together with pRL-SV40 (0.1 μg; Promega, Madison, WI) carrying the Renilla luciferase gene under the control of the cytomegalovirus promoter. Luciferase activity was measured using a luciferase assay system (Promega). The reporter activity was calculated by normalizing the firefly luciferase value with that of the Renilla luciferase control vector and expressed as relative luciferase units. All data were obtained from at least three independent experiments. The relative promoter activities were depicted as the mean ± S.D.
Immunofluorescence was performed on cultured WT and Nuc1 cells and retinal flat mounts as described 55. Fluorescent images were taken with LSM 510 confocal microscope.
WT and Nuc1 astrocytes in 6 well plates were transfected with scrambled, Cryba1, or Jagged1 ON-TARGETplus SMARTpool siRNA (Dharmacon, Lafayette, CO).
The Notch expression plasmid NICD was generated by PCR from full length Notch1 cDNA, corresponding to amino acids 1760 to 2556, and was cloned into pcDNA3.1 (Invitrogen). The NδE plasmid encodes the transmembrane and entire intracellular domain of murine Notch1, extending from I1701 to K2530 plus the C-terminal Myc tag in pcDNA3.1 mammalian expression vector (Invitrogen). The constructs NδE, NδE(V1744K), and ICv1744 were kindly provided by Dr. Raphael Kopan and have been described 56. Jagged1 overexpression plasmid containing a full length rat Jagged1 was cloned into pCDNA3. Jagged1 EGF repeats were deleted using site directed mutagenesis. Full-length rat Notch1 cDNA (Notch-FL), containing a single internal Myc epitope tag cloned into the pcDNA3.1. NEXT plasmid was made from the Notch-FL plasmid and NICD plasmid was purchased from Addgene. A GFAP-promoter reporter vector containing 1.9kb of the rat GFAP promoter upstream of the firefly cDNA was cloned into pGL2 vector as previously described 35.
The luminal pH of endolysosomal vesicles was measured by a ratiometric method using a fluid phase fluorescent marker, fluorescein isothiocyanate (FITC)-conjugated dextran (Invitrogen) 57. Cells were incubated with FITC-dextran (500 μg/ml, Invitrogen) for 3 hours to allow internalization of the fluorescent probe, then washed and resuspended in 50 mM MES buffer (pH 6.5), and the fluorescence measured at 520 nm by exciting at 450 nm and 495 nm. The fluorescence excitation ratio (I495nm/I450nm) was calculated, and endosomal pH determined from a calibration curve of FITC-dextran in the pH range of 4 to 7. To generate the calibration curve, cells loaded with FITC-dextran were harvested and resuspended in 10 mM phosphate buffer supplemented with 2 Mm FITC-dextran at 2 mg/ml and the measurements were made over time 58.
For the measurement of lysosomal V-ATPase activity, acridine orange uptake assay was performed with isolated lysosomes 59. The lysosomes were suspended in buffer containing 6 μM acridine orange (Invitrogen), 150 mM KCl, 2 mM MgCl2, and 10 mM Bis-Tris-propane. After a steady spectrofluorometric baseline was reached, V-ATPase was activated by addition of ATP (1.4 μM final concentration) and 2.5 μM valinomycin at pH 7.0 to promote the movement of K+ from the inside to the outside of the lysosome, generating a membrane potential. Acridine orange uptake is a measure of V-ATPase-driven pumping of hydrogen ions into lysosomes. Fluorescence at 530 nm (excitation at 495 nm) was measured using a fluorescence plate reader (Bio-Tek Synergy, Winooski, VT). Separately, isolated lysosomes containing the same activation buffer were used to measure the extralysosomal quenching of acridine orange fluorescence. To confirm V-ATPase activity, stimulation of intralysosomal fluorescence and quenching of extralysosomal fluorescence were measured with or without 1 μM bafilomycin A1 (Sigma).
To prepare endosomal fractions, cells were homogenized in 1.5 ml of 2 M sucrose in HEPES-EDTA buffer containing protease and phosphatase inhibitor cocktails (Sigma). A postnuclear supernatant (PNS) was prepared by centrifugation at 1,000 × g at 4°C for 10 min to remove the cell debris, followed by 3,000×g for 10 min to remove the nuclei. PNS (0.5 ml) was diluted 1:1 with sucrose (40%) and imidazole (3 mM) solution in HEPES-EDTA buffer. After overlay with 1.5 ml of 35% sucrose with 3 mM imidazole, 1 ml of 25% sucrose with 3 mM imidazole, and 1 ml 8% sucrose with 3 mM imidazole, the sucrose gradients were centrifuged in a SW55 rotor (Beckman, Indianapolis IN) at 150,000 × g for 90 min. Translucent interfaces were collected from the top of the centrifuge tube.
Cell homogenates were centrifuged at 3000×g for 10 min to remove nuclei and at 10, 000×g for 15 min to remove mitochondria. This supernatant was subjected to density gradient centrifugation and sample collection as described 32.
The iodixanol fractions were incubated with 20 mM Hepes, 150 mM NaCl, 5 mM EDTA, pH 7.0) and 0.4% CHAPSO and a protease inhibitor cocktail (Sigma) at 37°C for 16 h with or without 1 μM of the γ-secretase inhibitor L-685,458 (Bachem, Torrance, CA). Reactions were stopped by addition of RIPA buffer, heated to 95°C for 5 min and centrifuged at 16 000×g for 5 min. Production of Aβ40 was analyzed by ELISA with anti-Aβ40 antibody according to the manufacturer’s protocol (Wako chemicals, Richmond, VA). Production of Aβ40 was calculated as Aβ40 levels without L-685,458 minus Aβ40 levels with added L-685,458. Cells incubated with the lysosomal inhibitor, choloroquine were also used to assay the activity of γ-secretase.
Statistical analysis was performed using Microsoft excel and GraphPad 5.0 software. The P-values were determined by two-tailed Student’s t-test in a triplicate experiment representative of at least three independent experiments. Multiple comparisons were made using Kruskal-Wallis test. Significance was defined as *P≤0.05, *P≤0.01. Results are presented as mean+ SD. Each biological replicate has at least 5 technical replicates.
This work was supported by grants from the National Institutes of Health, EY018636 (DS), EY019037 (DS) and EY01765 (Wilmer Imaging Core) and Research to Prevent Blindness (an unrestricted grant to The Wilmer Eye Institute). DS is a recipient of the Sybil B. Harrington Special Scholar award for Macular Degeneration from Research to Prevent Blindness. We would like to thank Drs. Akrit Sodhi and Xiaoban Xin for providing the adenoviral vectors and Dr. Raphael Kopan for NδE plasmids. We also thank Morton F Goldberg, Bidyut Ghosh and Seth Greenbaum for critically reading the manuscript.
AUTHOR CONTRIBUTIONSMV conducted the experiments, participated in the study design and analyzed data. SH constructed figures for the manuscript and performed some initial experiments. CG contributed to the transcriptional studies. LD and EFW participated in the generation of Cryba1 floxed mice. SZ participated in the study design and analyzed data. DS designed the study and assisted with the generation of Cryba1 floxed mice and data analysis. MV, SZ and DS wrote the paper. All authors have approved the final manuscript.
COMPETING FINANCIAL INTERESTS
The authors declare no competing financial interests.