|Home | About | Journals | Submit | Contact Us | Français|
We have previously shown that myelin abnormalities and loss characterize the normal aging process of the brain and that an age-associated reduction in Klotho is conserved across species. Predominantly generated in brain and kidney, Klotho overexpression extends life span, whereas loss of Klotho accelerates the development of aging-like phenotypes. While the function of Klotho in brain is unknown, loss of Klotho expression leads to cognitive deficits. In the present study, we found significant effects of Klotho on oligodendrocyte functions including induced maturation of rat primary oligodendrocytic progenitor cells (OPCs) in vitro and myelination. Phosphoprotein Western analysis indicated Klotho's downstream effects involve Akt and ERK signal pathways. Klotho increased OPCs maturation, and inhibition of Akt or ERK function blocked this effect on OPCs. In vivo studies of Klotho knockout mice and their control littermates revealed that knockout mice have a significant reduction in major myelin protein and gene expression. By immunohistochemistry, the number of total and mature oligodendrocytes was significantly lower in Klotho knockout mice. Strikingly, at the ultrastructural level, Klotho knockout mice exhibited significantly impaired myelination of the optic nerve and corpus callosum. These mice also displayed severe abnormalities at the nodes of Ranvier. In order to decipher the mechanisms by which Klotho affects oligodendrocytes, we used luciferase pathway reporters to identify the transcription factors involved. Taken together, these studies provide novel evidence for Klotho as a key player in myelin biology, which may thus be a useful therapeutic target in efforts to protect brain myelin against age-dependent changes.
Klotho is an anti-aging protein named after the mythical Greek goddess who “spins the thread of life” (Kuro-o et al., 1997). Klotho knockout mice exhibit many changes that also frequently occur during human aging, including arteriosclerosis, osteoporosis, and cognitive decline. The mice develop normally but die prematurely, with an average lifespan of ~61 days (Kuro-o et al., 1997), whereas mice overexpressing Klotho live 30% longer than wild type mice (Kurosu et al., 2005). In early adulthood, Klotho knockout mice show memory retention deficits (Nagai et al., 2003b), a reduction in synapses in the hippocampus (Li et al., 2004), perturbations in axonal transport, and a neurodegenerative phenotype in the hippocampus (Shiozaki et al., 2008). Interestingly, humans carrying the Klotho-VS polymorphism exhibit reduced cognitive abilities (Deary et al., 2005; Nagai et al., 2003b), as well as reduced life span (Arking et al., 2005; Arking et al., 2003; Nagai et al., 2000). The basis of cognitive deficits in Klotho knockout mice, or in humans with Klotho-VS polymorphism is not known, and a more thorough assessment of neuropathology is needed.
Klotho is highly expressed in the choroid plexus and neurons, especially in the hippocampus, as well as in the kidney and reproductive organs (Kuro-o et al., 1997), but its function in the brain is not clear. Klotho is a type I transmembrane protein cleaved by ADAM10 and 17 from the cell membrane (Bloch et al., 2009; Chen et al., 2007), and is detectable in serum and CSF (Imura et al., 2004). Klotho functions include regulation of FGF23 signaling, suppression of the insulin/IGF1 and Wnt signaling pathways, and regulation of calcium and phosphate homeostasis (Kuro-o, 2010). In contrast to other organ systems, none of the downstream effects of Klotho have been studied for the CNS and little is known about downstream signal transduction machinery required for Klotho's CNS effects. Considering the distribution of Klotho expression and processing within the CNS and the effects on cognition cited above, Klotho is positioned to influence a variety of CNS structures and functions in development and aging.
As part of our studies of age-associated cognitive decline in the rhesus monkey, we discovered using microarray analysis that Klotho expression was decreased in the aged corpus callosum (Duce et al., 2008), likely due to the hypermethylation of its promoter (King et al., 2011). This is of interest as damage to myelin is ubiquitous in aging monkey brain (Kohama et al., 2011) at biochemical (Sloane et al., 2003), ultrastructural (Bowley et al., 2010) and macroscopic levels (Makris et al., 2007; Wisco et al., 2008) and these abnormalities are often associated with cognitive decline (Hinman and Abraham, 2007; Peters, 2009). To determine whether there is a connection between age-related alterations in Klotho expression (Duce et al., 2008) and CNS dysmyelination (Hinman and Abraham, 2007; Sloane et al., 2003), and to understand the functions of Klotho in the brain, we assessed how Klotho influences oligodendrocyte function and developmental myelination and describe here the important and novel role Klotho plays in oligodendrocyte biology and myelination.
The recombinant mouse Klotho protein containing the extracellular domain of mouse Klotho (Ala 35- Lys 982) was from R&D Systems (Minneapolis, MN). Akt inhibitor LY294002 and ERK inhibitor UO126 were from Cell Signaling (Danvers, MA). Growth factors for oligodendrocytes were from Peprotech (Rocky Hill, NJ). All other chemicals were from Sigma-Aldrich (St. Louis, MO).
Klotho knockout (Kuro-o et al., 1997) and control mice were perfused with 0.1 M Phosphate buffer saline pH 7.4 at 4°C and hemibrains homogenized in 5× (v/w) ice-cold RIPA buffer with protease and phosphatase inhibitor cocktails (Roche). Samples were then centrifuged at 16,000×g for 15 min, and the supernatant was collected for SDS-PAGE and Western blot analysis.
OPCs were isolated from female Sprague-Dawley postnatal day 2 rat pups as described previously (Mi et al., 2005; Sloane and Vartanian, 2007). Cultures were maintained in high-glucose DMEM OPC culture medium (4 mM L-glutamine, 1 mM sodium pyruvate, 0.1% BSA, 50 μg/mL insulin, 30 nM sodium selenite, 10 nM D-biotin and 10 nM hydrocortisone) containing bFGF/PDGF (10 ng/mL) for 1–3 days and replaced with either DMEM medium with CNTF (10 ng/mL), T3 (15 mM) and NT3 (10 ng/mL), or DMEM with CNTF and NT3 alone, with or without recombinant mouse Klotho at a concentration of 0.4 μg/mL at 37°C for 3–8 days, as indicated in the figures. Half of the medium bathing the cells was replaced with fresh medium with or without Klotho every other day. The same concentration of Klotho was used in all experiments. The cell lysates were collected in RIPA buffer as described before, and protein samples were analyzed by SDS-PAGE and Western Blotting.
Protein concentrations were measured using the Micro BCA Protein Assay Reagent Kit (Pierce, Rockford, IL) according to the manufacturer's protocol. For SDS-PAGE, cell lysates containing the same amount of total protein were boiled for 5 min and loaded on 4–20% pre-cast Tris-Glycine gels (Bio-Rad, Hercules, CA). Proteins were transferred to nitrocellulose membranes (Millipore, Billerica, MA). All antibodies were diluted in TBST (50 mM Tris, pH 8.0, 150 mM NaCl and 0.1% (v/v) Tween 20) containing 5% (w/v) nonfat dry milk. Secondary antibodies were horseradish peroxidase-conjugated goat anti-mouse, anti-rat or anti-rabbit (1:5000, Kirkegaard & Perry Laboratories, Gaithersburg, MD). Enhanced chemiluminescence (ECL) was detected using Immobilon Western Chemiluminescent Substrate (Millipore, Billerica, MA). Autoradiography was done using Kodak Scientific Imaging Film X-OMAT™ AR (Eastman Kodak, Rochester, NY).
The primary antibodies used were: anti-MBP mouse monoclonal antibody (1:1000, Covance), anti-MAG mouse monoclonal antibody (clone B11F7, 1:1000), anti-CNP mouse monoclonal antibody (1:1000, Sigma, St. Louis, MO), anti-FGF receptor substrate (FRS) Y196 (1:1000, Cell Signaling, Danvers, MA), anti-OSP rabbit polyclonal antibody (1:1000, Abcam), anti-PLP mouse monoclonal antibody (1:1000, Millipore), anti-PCNA clone PC10 (1:1000, Upstate, Temecula, CA) and anti-β-Tubulin monoclonal antibody (1:1000, Invitrogen, Carlsbad, CA). The antibodies in the Akt and ERK pathways were from phospho Akt and ERK pathway kit (Cell Signaling, Danvers, MA) and were used according to the manufacturer's protocol.
Total RNA was isolated using QIAGEN's RNeasy kit (Qiagen, Valencia, CA). A reverse transcription was performed with 2 μg of total RNA from each sample. Primers for selected mouse myelin related genes and corresponding rat oligodendrocyte maturation enriched genes as published (Cahoy et al., 2008) were designed by RealTimePrimers. Controls included beta actin (ACTB), beta-2 microglobulin (B2M), Phosphoglycerate kinase 1, hypoxanthine phosphoribosyltransferase 1 (Hprt1) and Glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The qRT-PCR experiments were performed with iQ™ SYBR® Green Supermix (Bio-rad) with detection on a Bio-rad C1000 Thermal Cycler. The whole cDNA from 2 μg of total RNA was used for one 96 well plate. Triplicates of 20 μL reactions containing primer and cDNA template were used for quantitation. A PCR reaction was carried out as follows: 1 cycle of 95°C for 3 min followed by 40 cycles of 95°C for 10 s, 55°C for 20 s, and 72°C for 30 s. This was followed by a dissociation curve beginning at 55°C and increasing by 0.2°C every 3 s, with SYBR green fluorescence measured at every interval. Relative quantitation of the difference between the control and Klotho-treated samples was done using RT2 Profiler PCR Array Data Analysis Program from Qiagen. Genes were tested for statistical significance (P < 0.05), relative to the control, by student t-test.
Pax3 reporter plasmid (pluc-TKCD-19) was kindly provided by Dr. Frank Rauscher III (Wistar Institute, Philadelphia, PA). Signal Transduction 45 pathway Reporter Array was from SABiosciences (Qiagen, Valencia, CA). OPCs were plated into 96 well plates at a density of 0.65×106 cells/plate in OPC culture medium containing bFGF/PDGF for 3 days before transfection. 200 ng of Luciferase reporter or control empty-vector DNA (with 5 ng Renilla luciferase) were transfected into cells using Lipofectamine 2000 (Invitrogen), and cells were treated with or without recombinant mouse Klotho in OPC culture medium as described above. Twenty-four hours post transfection, cells were washed once with PBS and assayed for luciferase activity using the Dual-luciferase system (Promega) as described (Oh et al., 2010).
Cells were fixed in 4% paraformaldehyde in PBS at room temperature, rinsed with PBS, and treated for 1 h with blocking solution (PBS supplemented with 0.1% Triton ×100 and 1% BSA). Cells were incubated overnight at 4°C with the primary antibody diluted in blocking solution. Cells were stained with antibodies to Olig2 (1:10,000) (Millipore), CC-1 (1:200) (Millipore), or O1 (1:2) (ATCC). Subsequently, cells were rinsed and incubated with the relevant secondary antibodies (Cy3 or Alexa-488) (Jackson ImmunoResearch) for 1 h at room temperature. Immunofluorescence images were obtained by a Nikon Eclipse 660 microscope and a SPOT-cooled CCD digital camera (Diagnostic Instruments). For every condition, we identified the maturation state of each OL for the ~30–60 cells within a given field for all fields acquired (N = 450–900 cells/condition) from 3 independent experiments. We quantified numbers of mature OLs (O1+ or CC1+) and total OL-lineage cells (Olig2+) and determined percent mature OLs of total OL-lineage cells (O1+/Olig2+ or CC1+/Olig2+) (since Olig2 labels OPCs as well as OLs).
Cell number was determined colorimetrically by crystal violet staining as described previously (Zeldich et al., 2007). Cells were plated at 50,000 cells/well in 24-well plates in triplicates, and allowed to attach and spread in OPC culture medium containing bFGF/PDGF for 1–3 days and then replaced with medium containing CNTF and NT3 with or without Klotho for the indicated time periods. At the end of each time interval, the cells were washed with PBS, fixed in 70% ethanol and kept at 4°C until the staining with 1% crystal violet. Unincorporated stain was removed by washing, cells were air-dried, and the dye was extracted with 70% ethanol and optical density (absorbance 550 nm and baseline reference absorbance 750 nm) was measured by Microplate Reader (Glomax Multi Detection System, Promega, USA). The data analysis was done by subtracting the baseline readings (750 nm) from the absorbance readings (560 nm).
For cell viability assay, cells were plated in 96-well plates in six repeats, and allowed to attach and spread in OPC culture medium containing bFGF/PDGF for 3 days and then replaced with medium containing CNTF and NT3 with or without Klotho for the indicated time periods. At the end of each time interval cell number was assessed using CellTiter-Glo Luminescent Cell Viability Assay (Promega, USA) according to the manufacturer's instructions. This assay signals the presence of metabolically active cells, Briefly, 100μl of CellTiter Glo Reagent was added to the equal volume of cell culture medium present in each cell culture well. The contents were mixed for two minutes and after stabilization of the signal at RT for 10 min, the luminescence was recorded on a Microplate Reader (Glomax Multi Detection System, Promega, USA). Cell death was assessed by using the CytoTox 96 non-radioactive cytotoxicity assay (Promega, USA), which measures the LDH release into the medium.
Five week old Klotho +/+ and −/− mice were perfused through the heart with 0.1M PBS (4°C, pH 7.4). The brains were removed and one hemisphere was immersion fixed in 4% paraformaldehyde overnight while the other was left unfixed and flash frozen with pulverized dry ice and stored at −80°C. The fixed hemisphere was removed from fixative the next day and cryoprotected in successive solutions of first 10%, then 20% glycerol in PBS with 2% DMSO (Rosene et al., 1986). The frozen hemibrains were cut in the coronal plane into 30μm thick sections with a sliding microtome. The sections were divided into six interrupted series and stored at 4°C in 0.1M PBS with 1% sodium azide as a preservative until they were processed.
For Olig2, CC-1 and GST-Pi immunohistochemistry, one series from each subject was selected and all sections were processed together in a single batch at the same time in the same reagents to eliminate any procedural variance. All steps were performed at RT unless otherwise indicated. Free-floating sections were washed, quenched with 3% hydrogen peroxide, and then blocked with 10% normal goat serum (NGS) in Phosphate Buffered Saline (PBS) with 0.4% Triton-X 100 for 1 hour. Next, sections were incubated in a stock solution (1% NGS, 0.2%Triton-X100 in 0.05M PBS) containing rabbit polyclonal anti-Olig2 antibody (Millipore ab9610, 1:10,000), mouse mAb against CC-1 (Millipore OP-80, 1:40) or anti-GST pi (Abcam, 1:500) for 48 hours at 4°C. Sections were then washed in stock solution and treated with biotinylated goat anti-rabbit or anti-mouse secondary antibody (1:600, BA5000, Vector Laboratories) for 1 hour. After washing, sections were incubated with an avidin-biotinylated horseradish peroxidase enzyme complex (PK6100, Vectastain Elite ABC kit, Vector Laboratories) for 1 hour, rinsed in KPBS, then sodium acetate buffer (0.175M), and processed with a solution of nickel sulfate hexahydrate (0.095M), 3,3 diaminobenzidine tetrahydrochloride (DAB) (0.55mM), and hydrogen peroxide (0.0025%) in 0.175M sodium acetate buffer for 15 min. The immunostained sections were mounted on gelatin-coated slides and air-dried. Then sections were defatted in chloroform:ethanol (1:1), rehydrated, counterstained with Neutral Red (1%), dehydrated in a series of ethanols, cleared in xylenes and coverslipped with Permount (Fisher Scientific).
To quantify Olig2+ and GST-Pi+ cells, the fractionator method was used to obtain a population estimate within the fimbria. The fimbria was chosen because it is an easily defined and circumscribed bundle of white matter. The fimbria was outlined in six 30 micron sections spaced 180 micron apart, starting rostrally at the level of the anterior end of the habenula and spanning approximately 1.08mm caudally using StereoInvestigator system (version 9.14.3, MBF Bioscience, Williston, VT) and a 4× Nikon Plan objective on a Nikon Eclipse E600 microscope. Cell counts were made with a 20× Nikon Plan Fluor objective. A 120 micron × 120 micron grid was overlaid on the region of interest and Olig2+ cells were counted within 40 micron × 40 micron counting frames placed at the grid intersections that fell within the outline of the fimbria. The criterion for counting cells was the appearance of the black Ni-DAB immuno-product in the nucleus. In an adjacent series of six 30 micron thick tissue sections, GST-Pi positive cells within the fimbria were quantified. The fimbria was outlined with a 4× objective as described above. A 120 micron × 120 micron sampling grid was superimposed over the region of interest and GSTPi positive cells were counted within a 40 micron × 40 micron counting frame and 5 micron disector using a 60× Nikon Plan Fluor oil objective. A guard volume of 1 μm above the disector box was used and 2–4 micron below, depending on the thickness of the tissue section (approximately 8–10 micron total). GST-Pi positive cells had brown DAB immuno-product in the cytoplasm and nucleolus, yet the nucleus was unstained.
Free floating 30 micron sections were stained for nodes and paranodes with rabbit anti-beta-IV spectrin (1:400) or rabbit anti-NaV1.6 (1:200) (gifts from M. Rasband, Baylor College of Medicine (Ogawa et al., 2006; Rasband et al., 2003), and mouse anti-caspr (Antibodies, Inc.) (1:500), using the following protocol with all steps performed at RT unless otherwise stated. The monoclonal antibody mouse anti-caspr (clone K65/35) was developed by and obtained from the UC Davis/NIH Neuromab Facility. Sections were washed in PBS, transferred to prewarmed 10 mM sodium citrate, pH 8.5 and incubated at 80°C for 30 min for antigen retrieval. After cooling to RT, sections were washed 3 times in PBS, then blocked and permeabilized in PBS with 0.3% TX-100 (Sigma) and 10% normal donkey serum (Jackson Immunoresearch) for 30 min. Sections were then placed directly in MOM Block (Vector Labs) diluted in PBS for 1 hr, washed 3 times in PBS and then incubated overnight in primary antibody cocktails. The next day, after 3 washes in PBS, sections were incubated in donkey anti-mouse Cy2 (Jackson Immunoresearch; 1:300) and donkey anti-rabbit DyLight 549 (Jackson Immunoresearch; 1:300) for 1 hr. Sections were then mounted on gelatin subbed slides, air dried and dehydrated with 70–100% EtOH and xylene and coverslipped with DPX mounting medium.
Image acquisition was performed on a Nikon Eclipse Ti C2+ laser scanning confocal microscope. Two-micron optical sections at 100× optical objective with 2× digital zoom were obtained from five different immunolabeled sections from each of two wild-type and two Klotho-null mice. The imaged white matter included corpus callosum overlying striatum to that overlying hippocampus. Maximum intensity projections were obtained and images were postprocessed in NIS Elements software (Nikon). Nodal and paranodal segments were measured using the two-point length measurement tool. For measurement, only complete paranodal pairs (both paranodes and an immunolabeled node) were selected for measurement. A total of thirty paranodes per section (150 per animal) were measured and a total of 10 nodes per section (50 per animal) were measured. Statistical significance was determined using a Student's t-test assuming one-tail and equal variance. For presentation, images were further post-processed and cropped in Photoshop CS5 (Adobe).
35 days old Klotho +/+, +/− and −/− mice, and 25 days old Klotho −/− mice were perfused transcardially with 1% paraformaldehyde and 2.5% glutaraldehyde fixative in 0.1 M cacodylate buffer pH 7.2–7.4. Optic nerve, corpus callosum and cervical spinal cord were postfixed with 1% osmium oxide, dehydrated in alcohol series and propylene oxide before embedding in Epon. One-micrometer-thick sections were cut and stained with toluidine blue. Thin sections were contrasted with uranyl acetate and lead citrate and imaged with a JEOL (Peabody, MA) 1200EX electron microscope.
GSEA was performed according to Subramanian et al. (Subramanian et al., 2005) with 1186 chemical and genetic perturbation (CGP) gene sets. Klotho −/− mice versus Wild-type +/+ mice gene expression profiles were ranked based on P value. The ranked gene list was utilized as input for the CGP gene sets. The significant gene sets were selected based on the normalized enrichment score (NES) and false discovery rate (FDR), accordingly.
Klotho has been reported to affect intracellular signaling through Akt and ERK1/2 pathways in HEK293 and breast cancer cells (Kurosu et al., 2006; Wolf et al., 2008). We therefore assessed whether Klotho induces ERK1/2 and Akt protein phosphorylation from 0 to 60 min in OPCs isolated from rat brain. Western blot results indicated that proteins in the ERK1/2 and Akt pathway including PTEN (Ser380), Akt (Ser473), and GSK-3β (Ser9) were phosphorylated upon Klotho treatment (Figure 1). We found two waves of GSK3βphosphorylation in OPCs upon Klotho treatment, with a first peak at 10 min and a second at 45 min (Figure 1). These results suggest that the Akt and ERK/12 pathways are both involved in Klotho-induced signaling in rat primary OPCs. We also examined phosphorylation of FGF receptor substrate 2 (FRS2) since Klotho may function by modulating FGF signal transduction pathways (Kurosu et al., 2006; Urakawa et al., 2006). Western blot results indicated that FRS2 was phosphorylated at Y196 upon Klotho treatment (Figure 1), suggesting that the FGF signaling pathways may be involved in Klotho-induced signaling in rat OPCs. However, it is also possible that Klotho affects FRS2 phosphorylation via Klotho's interaction with the TrkA receptor (Meakin et al., 1999).
To examine the effects of Klotho on the OPC phenotype, oligodendrocyte maturation was assessed by immunohistochemistry for mature oligodendrocytes (O1), and pan-oligodendrocytes (Olig2) after Klotho treatment. The OPCs were allowed to attach and spread in OPC culture medium containing bFGF/PDGF for 3 days and then replaced with medium containing CNTF and NT3 with or without Klotho. Following 3 and 6 day Klotho treatment, we found that Klotho increased the percentage of mature primary oligodendrocytes (% O1/Olig2) (Figure 2A and B). Similar results were obtained using another marker of mature oligodendrocytes, CC-1, (Figure 2C). Klotho enhances OPC maturation in medium containing CNTF and T3, or in medium containing CNTF, T3 and NT3 (Figure 2D). Since both Akt and ERK1/2 phosphorylation occurred rapidly after exposure to Klotho, we asked whether Akt and ERK1/2 function is required for Klotho's effects on OPC maturation. OPCs were treated with Klotho in the presence or absence of Akt and ERK inhibitors (LY294002 (LY) for Akt inhibition or UO126 (UO) for ERK inhibition). Inhibition of ERK function reduced, while Akt functional inhibition completely abolished the effects of Klotho on OPC maturation (Figure 2E), suggesting that Klotho enhances OPC maturation primarily via signaling requiring Akt but also, to a more limited degree, through ERK1/2. Western blot analysis of Klotho-treated OPCs revealed that Klotho enhanced the expression of the major myelin proteins including myelin-associated glycoprotein (MAG), myelin basic protein (MBP), oligodendrocyte specific protein (OSP/Claudin11), and proteolipid protein (PLP) (Figure 2F and G), confirming that Klotho enhances OPC maturation in vitro. Western blotting for the cell proliferation marker, PCNA, revealed no difference with Klotho treatment (Figure 4F and G), suggesting Klotho does not enhance OPC proliferation. We also examined cell number and cell viability by crystal violet staining and CellTiterGlo assay which reflects the amount of the ATP present, and we found no difference in Klotho treated and untreated OPCs from day 1 to day 4 in the cell viability assay and for day 2 and day 4 for crystal violet staining (Figure 2H and I). Since OPCs were allowed to attach and spread for 3 days in the presence of basic FGF (bFGF) and PDGF, cell proliferation occurred during this period. After we treated the OPCs with Klotho in culture medium containing CNTF and T3, OPCs started to differentiate but cell numbers remained constant as shown in Figure 2H and I. We also observed no difference with a cell death assay by measuring LDH release to the media with or without Klotho. These results suggest that Klotho affects oligodendrocyte differentiation and maturation, but not their proliferation and cell death.
Recently a transcriptome database for astrocytes, neurons and oligodendrocytes has been published providing cell type specific markers for these neural cells (Cahoy et al., 2008). To confirm the effect of Klotho on OPC maturation, we performed qRT-PCR analysis of the top 45 genes (see Table 4 for primers information) from the OL maturation enriched gene list (Cahoy et al., 2008) using RNA isolated from OPCs treated with Klotho or PBS only, for 3 or 7 days. qRT-PCR revealed that comparing 3 days and 7 days differentiated OPC, most of the genes were highly expressed at day 7 but had either low or undetectable expression at day 3 (Figure 3A), suggesting the primer sets were working well to detect OPC maturation enriched genes. We then analyzed the gene expression changes of RNA from OPC treated with Klotho for 7 days compared to PBS-treated control. We found that 37 out of the 45 genes were detectable, and 78% (29/37) of the genes were up-regulated by Klotho. Of those 37, 21 genes reached significance of p<0.05 (Figure 3B). The qPCR results were consistent with the results obtained from protein expression as assessed by WB in that the expression of major myelin proteins was enhanced 2–3 folds in response to exogenous addition of Klotho (Figure 2F and G). These results demonstrate that Klotho can enhance OPC maturation as shown by the immunostaining of the cells with two separate oligodendrocyte markers O1 and CC-1, and by WB analyses as shown in Figures 2B, D, F and G.
If Klotho is important for oligodendrocyte maturation in vitro, Klotho knockout mice should exhibit deficiencies in myelination. The ultrastructure of 35-day-old Klotho knockout (−/−), hemizygous (+/−) and control (−/−) mice brains was examined by electron microscopy. There was a significant reduction in the percentage of myelinated fibers in both optic nerve and corpus callosum, but not cervical spinal cord (Figure 4), suggestive of region-specific effects of Klotho expression. In the optic nerve, Klotho −/− mice exhibited a drastic reduction in the percentage of myelinated fibers (10% compared to 90% in +/+ littermates) (Figure 4E, F and M), while Klotho +/− mice showed normal numbers of myelinated fibers (Figure 5C and D compared to A and B). In the corpus callosum, both Klotho +/− and Klotho −/− exhibited significantly impaired myelination (20% compared to 90% in +/+ mice) (Figure 4G–M). No significant change was seen in the percentage of myelinated fibers in the spinal cord (Figure 4M). A similar reduction in the myelination of fibers in the corpus callosum was found in two 25 day-old Klotho knockout brains, with only 16.4±1.6% fibers being myelinated.
In order to decipher the potential basis for region-specific effects of Klotho, we examined the distribution of Klotho in the corpus callosum, spinal cord and optic nerve from five 1.1 and five 1.6 month-old mice. The results showed that Klotho was expressed in all three regions, with the highest expression in the corpus callosum, and lowest in spinal cord at the younger age (1.1 months, Figure 4N and O). The results of the WBs of the three specific brain samples represent the levels of available Klotho protein surrounding these three brain regions. A higher Klotho level, e.g, corpus callosum, implies that Klotho's function is needed in that region, and less Klotho suggests Klotho is less necessary, e.g, spinal cord. These data, together with the EM results, suggest that Klotho is required for proper myelination of fibers in the corpus callosum. We therefore examined the myelin markers myelin associated glycoprotein (MAG), MBP, and 2', 3'cyclic nucleotide-3'-phosphodiesterase (CNP) in brain tissues of 8 week old Klotho knockout and control mice. Western blot analyses demonstrated significant reduction of major myelin proteins in Klotho deficient mice compared to control mice (Figure 4P and Q) indicating that Klotho affects myelination via inducing myelin protein production by oligodendrocytes.
To assess the effects of the hypomyelination seen in Klotho −/− mice on axonal microdomain organization, free-floating sections were stained for caspr to label paranodes, and beta-IV spectrin or Nav1.6 to label nodes. No differences were seen in nodal staining patterns between beta-IV spectrin and Nav1.6 immunolabels. High magnification confocal optical sections reveal that within the corpus callosum, Klotho −/− mice show a significant decrease in paranodal length compared to wild-type controls (Figure 5). This decrease in paranodal length suggests that there is less robust myelination in each axon in the Klotho −/− mice consistent with hypomyelination seen in the ultrastructural analysis. While the absolute difference in paranodal length is small (0.30 microns), this average value would translate to one less paranodal loop per axon (assuming each loop is roughly 0.25 microns, (Trapp and Kidd, 2004)). The decrease in paranodal length was associated with a corresponding significant increase in nodal length by 0.25 microns (Figure 5E). Analysis of nodal and paranodal domains in the corpus callosum (Figure 5F) and optic nerve (data not shown) by electron microscopy did not demonstrate any ultrastructural changes in axonal microdomains. Since nodal length shortens during myelination (Dugandzija-Novakovic et al., 1995; Vabnick et al., 1996), this increase in nodal length likely represents more immature nodal segments in Klotho −/− mice. Longer nodes may be more energetically unstable and lead to progressive axonal degeneration (Waxman, 1998).
Immunohistochemistry and quantitative analysis of brain sections from Klotho knockout mice at 5 weeks of age showed a ~30% reduction in the numbers of Olig2+ oligodendrocyte-lineage cells in the fimbria compared to control mice (Figure 6A–E). In the corpus callosum, we detected a similar trend towards a significant reduction in Olig2+ cells in Klotho knockout mice (data not shown). In addition, we found a similar and confirmatory reduction (21%) in the number of mature oligodendrocytes immuno-positive for the GST-Pi marker (Mason et al., 2004), suggestive of reduced levels of GST-Pi-positive mature oligodendrocytes in the fimbria of Klotho null mice compared to control mice (Figure 6F–J).
In order to determine whether these changes in the expression of myelin proteins are also seen at the mRNA level, we performed microarray analysis using RNA from 8 weeks old Klotho +/+ and −/− mice brain tissue, and applied gene set enrichment analysis to identify which canonical pathways were involved. With this approach, we found that a set of myelin related genes were downregulated. To examine whether Klotho affects gene expression of these myelin related genes, we performed qRT-PCR analysis of the 15 myelin-related genes. The qRT-PCR results revealed that all of the myelin related genes tested were reduced compared to control mice, and 11 out of 15 genes were down-regulated more than 2 fold (Table 1). The results demonstrate that lack of Klotho resulted in reduction in expression of major myelin genes.
To identify potential transcription factors involved in Klotho-mediated responses in OPCs, we examined regulatory effects of Klotho using a signaling pathway luciferase reporter system. We optimized transfection efficiency in primary oligodendrocytic cells by testing 15 available transfection reagents and found that Lipofectamine 2000 (Invitrogen) was superior for the luciferase reporter system in these cells (data not shown). We then tested 45 signaling pathway reporters (Qiagen, see Table 2) in OPCs with or without Klotho treatment. In addition, we also tested the Pax3 reporter since Pax3 repression is directly relevant to oligodendrocyte maturation to a myelinating phenotype (He et al., 2007). The results showed that Klotho inhibited C/EBP, AP1, NFkB, and Pax3 activities, while it enhanced LXR, ARE, STAT3, PR, and SRE MAPK/ERK transcription factor activities (Figure 7B, Table 3). The effect of Klotho on STAT3 phosphorylation in OPCs was also confirmed by Western blot (Figure 7A). These results revealed the possible transcription factors involvement in Klotho-mediated responses in OPCs.
In prior work, our group discovered reductions in Klotho in the aging brain white matter (Duce et al., 2008), where myelin deficits are also observed (Hinman and Abraham, 2007), suggesting a connection between Klotho expression and myelination. Thus, in this manuscript, we focused on the effect of Klotho on oligodendrocytes, the myelinating cells of the CNS. Our findings indicate a dramatic and important effect of Klotho on oligodendrocyte maturation and on myelination. Klotho enhances expression of MBP, MAG, PLP and OSP/claudin11, all being major myelin proteins, and induces OPC maturation in vitro. In vivo, Klotho knockout mice suffer from severe hypomyelination of the optic nerve and corpus callosum, which alters axonal microdomain organization. Myelin defects associated with loss of Klotho expression strongly indicate that decreased Klotho expression could result in slowed action potential propagation, conduction block, and destabilized coordinated neuronal communication, as seen in hypomyelinating and demyelinating diseases (Hanafy and Sloane, 2011; Hinman and Abraham, 2007). The numbers of myelinated fibers in the optic nerves and corpus callosum of the Klotho knockout mice were much lower that in the wild type mice, as determined by EM studies (Figure 4). However, the numbers of mature and total oligodendrocytes in the fimbria were reduced by a lower percentage (Figure 6). These data suggest that those oligodendrocytes that appear mature as judged by their staining with GST-Pi, were not able to myelinate axons properly, indicating an important role of Klotho in the myelination process.
We found that Akt and ERK1/2 signaling likely regulate OPC cells subjected to Klotho treatment. This finding is consistent with the data reported by Wolf et al. who found that both pathways were involved in Klotho-treated HEK293 and breast cancer cells (Wolf et al., 2008). Moreover, a recent study reported that Klotho induced ERK/MEK phosphorylation in HUVEC cells (Maekawa et al., 2011). Our results also suggest that Klotho could act on an as-yet unidentified Klotho receptor expressed on oligodendrocytes.
Oligodendrocyte development is regulated by numerous growth factors, which can signal through Akt and ERK1/2 dependent pathways (Bansal et al., 2003; Baron et al., 2000; Bhat and Zhang, 1996; Bibollet-Bahena et al., 2009; Cui and Almazan, 2007; Du et al., 2006; Fortin et al., 2005; Frederick et al., 2007; Frost et al., 2009; Fyffe-Maricich et al., 2011; Guardiola-Diaz et al., 2012; Van't Veer et al., 2009; Younes-Rapozo et al., 2009). Klotho has been reported to regulate insulin/IGF-1, FGF23, and Wnt signaling (Kuro-o, 2006, 2008a). In OPCs, Klotho is unlikely to function through IGF-1 signaling since Klotho inhibits the insulin/IGF-1 signaling pathway (Kurosu et al., 2005) and IGF-1 was reported to enhance OPC maturation and myelination (Bibollet-Bahena and Almazan, 2009). It is unknown whether FGF23 for which Klotho serves as a co-receptor in the kidney, has a role in neurodevelopment. However, our findings show that Klotho treatment of OPCs induces the phosphorylation of FRS2, a positive regulator of FGF signaling upstream of Akt and Erk pathway which is phosphorylated in oligodendrocytes upon activation of FGF-receptor (Bryant et al., 2009). Another putative receptor on OPCs is the NGF receptor, TrkA, since FRS2 is able to bind directly to both FGF and NGF receptors (Meakin et al., 1999; Ong et al., 2000). Possibly, Klotho functions through the FGF signaling pathway indirectly via another FGF ligand, or Klotho could modulate signaling of other cytokines in astrocytes via FGF signaling, as has been shown for insulin/IGF-1 signaling in C. elegans (Chateau et al., 2010). We cannot rule out the possibility that Klotho sequesters a ligand and blocks its accessibility to the receptor, or that Klotho binds directly to a receptor to prevent a ligand-receptor interaction. Another reported mechanism for Klotho effects is via the inhibition of the Wnt/β-catenin pathway, where Klotho may function as a secreted Wnt antagonist (Liu et al., 2007). Activating the Wnt/β-catenin pathway delays the development of myelinating oligodendrocytes (Fancy et al., 2009; Feigenson et al., 2009), while Olig2-induced neuronal stem cell differentiation into mature oligodendrocytes involves downregulation of the Wnt signaling pathway (Ahn et al., 2008). Whether Klotho stimulates oligodendrocyte maturation through the Wnt or insulin/IGF signaling pathways in addition to the FGF and NGF pathways is currently under investigation. It is interesting to note that in vivo, FGFR1/2 and ERK1/2 MAPK signaling in oligodendrocytes has recently been shown to regulate the growth of the myelin sheath, independent of oligodendrocyte differentiation (Furusho et al., 2012; Ishii et al., 2012).
We identified a set of transcription factors (TF) potentially involved in Klotho-dependent effects on OPC cells using the luciferase reporter system. Several of the TFs have been reported previously to be associated with Klotho functions, including C/EBP, AP1/JNK, NFkB, STAT3, and SRE MAPK/ERK (Chihara et al., 2006; Hsieh et al., 2010; Liu et al., 2011; Medici et al., 2008; Thurston et al., 2010; Wolf et al., 2008; Yamamoto et al., 2005; Zhao et al., 2011). Klotho protein increases resistance to oxidative stress at the cellular and organismal level in mammals (Kurosu et al., 2005; Nagai et al., 2003b; Rakugi et al., 2007; Shih and Yen, 2007). Klotho-deficient mice have impaired cognitive function when compared with wild-type mice, and treatment with the antioxidant α-tocopherol improves cognition (Nagai et al., 2003a), suggesting that Klotho may function as a neuroprotective factor. In addition, Klotho protein showed a neuroprotective effect on human neural stem cells (Foster et al., 2011), likely through the anti-oxidative stress effect of Klotho. We found an increase in the antioxidant response element (ARE) activity, which indicates that the antioxidative function of Klotho may be involved in the protection of oligodendrocytes. The new TF activities identified to be associated with Klotho include Pax3, LXR, and the progesterone receptor (PR). LXRs are oxysterol and nuclear receptors, which play an important role in the control of cholesterol homeostasis (Repa and Mangelsdorf, 2000; Whitney et al., 2002).
The lack of LXRs in mice leads to disorganized myelin sheaths, suggesting a role for the LXRs in myelination (Wang et al., 2002). Paired homeodomain protein 3 (Pax3) is important for development of myelinating glia and the myelination process (Wegner, 2000). An inverse correlation was observed between expression of Pax3 and MBP, suggesting that it represses MBP transcription (Kioussi et al., 1995). We found that Klotho inhibited Pax3 reporter activity and increased MBP expression in OPCs, therefore, we speculate that Klotho may increase MBP through the Pax3 pathway. Further investigation is required to decipher the direct or indirect involvement of the Pax3 transcription factor pathway resulting in Klotho's effects on oligodendrocytes. The transcription factor STAT3 has been previously reported to be involved in oligodendrocyte differentiation (Dell'Albani et al., 1998; Massa et al., 2000). Klotho treatment induced STAT3 phosphorylation, which is required for STAT3 activity (Bromberg and Darnell, 2000). Furthermore, the protein tyrosine phosphatase SHP-1 regulates oligodendrocyte differentiation through STAT3 in response to the IL-6 family of cytokines (Massa et al., 2000). Thus, Klotho may also act through STAT3 to drive OPC maturation.
Regardless of which receptor, signaling pathway(s), and transcription factors are involved, our in vitro and in vivo studies indicate that Klotho clearly has a significant regulatory role in OPC maturation and developmental myelination. Interestingly, Klotho has been reported to have a role in cell differentiation of diverse systems throughout the body including bone, fat and the cardiovascular system (Chihara et al., 2006; Kawaguchi et al., 1999; Shimada et al., 2004). Klotho gene expression appears to cause the impairment of both osteoblast and osteoclast differentiation (Kawaguchi et al., 1999). In addition, both angiogenesis and vasculogenesis are impaired in Klotho mutant mice, suggesting a role for Klotho in differentiation of endothelial precursor cells (Shimada et al., 2004). As a humoral factor, Klotho works to promote expression of differentiation markers in 3T3-L1 cells, indicating Klotho may play an essential role in adipocyte differentiation (Chihara et al., 2006).
In summary, we demonstrate a novel function of Klotho in oligodendrocyte maturation and developmental myelination of the CNS. This role is in addition to Klotho's function as a neuroprotective factor through preventing neurons from oxidative damage as proposed (Kuro-o, 2008b). Klotho may function as a humoral factor secreted by neurons or choroid plexus to promote myelination in neurodevelopment. It is possible that Klotho plays a regulatory role in maintaining or supporting oligodendrocyte and OPC function in the adult CNS, once the development has plateaued. Since we observed downregulation of Klotho in aged brain white matter, it is plausible that reduced Klotho level may account for damage to myelin and age-associated cognitive decline, and increasing Klotho level may protect myelin integrity and prevent myelin degeneration in the aged brain. Klotho is thus a new member of the large family of proteins that are crucial to neuron-oligodendrocyte communication, and studies on the functions of Klotho are likely to provide new therapeutic approaches for diseases in which myelin abnormalities play important pathogenic roles such as multiple sclerosis and schizophrenia (Edgar et al., 2004; Taveggia et al.).
We are grateful to Dr. Alan Peters for help in interpretation of the EM data. We thank Drs. Yuriy Alekseyev and Marc Lenburg for help with the microarray analysis and Drs. Sha Mi, Paul Rosenberg and Ms. Jianlin Wang for help with the early rat OPC preparations. We thank Dr. Cindy Lemere and Mr. Jeffrey Frost for the mice samples and help in isolating mouse brain tissues, Dr. Rong Fan for help in preparing Klotho mice brain tissue homogenates, and Ms. Chun-Tsin Hsu for assistance with the qRT-PCR experiments. We thank Dr. Matt Rasband for sharing of antibody reagents. We thank Drs. Gwendalyn King, James J. Collins, Ji-Hye Paik, Ido Wolf and Tami Rubinek for critical review of and suggestions for the manuscript, This work was supported by NIH-NIA grant AG-00001, an ADDF award to CRA, a NIH-NINDS R25 Training Grant to JDH, a National Multiple Sclerosis Society Research Grant to JAS, and an Ellison Foundation Award to CC.