Significance Global depletion of klotho accelerates aging, whereas klotho overexpression counteracts aging-related impairments. Why klotho is expressed at much higher levels in the choroid plexus than in other brain regions is unknown. We demonstrate in mice that aging is associated with klotho depletion in the choroid plexus. Reducing klotho selectively within the choroid plexus triggered inflammation within this structure and enhanced activation of innate immune cells within an adjacent brain region following a peripheral immune challenge. In cell culture, we identified a signaling pathway by which klotho suppresses activation of macrophages. Our findings shed light on klotho functions in the choroid plexus and provide a plausible mechanism by which klotho depletion from this structure promotes brain inflammation during the aging process.

Abstract Located within the brain’s ventricles, the choroid plexus produces cerebrospinal fluid and forms an important barrier between the central nervous system and the blood. For unknown reasons, the choroid plexus produces high levels of the protein klotho. Here, we show that these levels naturally decline with aging. Depleting klotho selectively from the choroid plexus via targeted viral vector-induced knockout in Klothoflox/flox mice increased the expression of multiple proinflammatory factors and triggered macrophage infiltration of this structure in young mice, simulating changes in unmanipulated old mice. Wild-type mice infected with the same Cre recombinase-expressing virus did not show such alterations. Experimental depletion of klotho from the choroid plexus enhanced microglial activation in the hippocampus after peripheral injection of mice with lipopolysaccharide. In primary cultures, klotho suppressed thioredoxin-interacting protein-dependent activation of the NLRP3 inflammasome in macrophages by enhancing fibroblast growth factor 23 signaling. We conclude that klotho functions as a gatekeeper at the interface between the brain and immune system in the choroid plexus. Klotho depletion in aging or disease may weaken this barrier and promote immune-mediated neuropathogenesis.

Aging is associated with a progressive increase in inflammatory alterations in the brain and other organs, a process that has been dubbed “inflammaging” (1). Biomarkers of inflammation are robust predictors of morbidity and mortality in older humans (1) and of age-related cognitive decline (2). The choroid plexus (CP) is an important gateway for the entry of immune cells into the central nervous system (CNS) (3, 4). Located within the ventricles of the brain, the CP consists of tight junction-bound epithelial cells resting upon a basal lamina and an inner stromal core. The stroma contains connective tissue and large capillaries with thin fenestrated endothelial walls and can harbor additional cell types, including dendritic cells with antigen-presenting capacity, macrophages, and granulocytes (3⇓–5). The CP responds to signals from blood and cerebrospinal fluid (CSF) and supports the CNS by producing CSF, nutrients, and growth factors and by regulating pH, osmolality, ion concentrations, and immune molecule content in the extracellular space of the brain and spinal cord (3⇓⇓⇓⇓–8). Throughout life, the CP produces much higher levels of the antiaging protein α-klotho (klotho) (9) than other components of the CNS [ref. 10 and the Allen Brain Atlas (portal.brain-map.org)]. However, the functions klotho fulfills in the CP remain to be defined.

Klotho production in peripheral organs is highest in the kidney where it regulates vitamin D metabolism and the transport of phosphate and calcium (8, 11). Alternative splicing gives rise to two isoforms: secreted klotho (s-KL, amino acids 1 to 550 in mice) and full-length transmembrane klotho (m-KL, amino acids 1 to 1014 in mice) (11). The latter can be cleaved by sheddases of the ADAM family, resulting in proteolytic release of klotho (p-KL, amino acids 1 to ∼960 in mice) into blood and CSF (11⇓–13). Klotho hypomorphic (kl/kl) mice have a deletion in the 5′ upstream region of the klotho gene, resulting in undetectable klotho mRNA levels in organs that normally express the klotho gene, such as the brain and kidney (9). While the possibility that the klotho gene is slightly transcribed in these mutants remains, the effect of this deletion is markedly reduced klotho expression throughout the body and a premature aging phenotype that affects multiple peripheral organs, resulting in early death (9, 14). CNS alterations in kl/kl mice include hypomyelination, increased expression or phosphorylation of neurofilaments, synaptic loss, and behavioral impairments (14⇓–16). Conceivably, at least some of these alterations result from the global reduction of klotho during early development and related systemic alterations. Whether more circumscribed reductions of klotho later in life also cause CNS pathology is not known. Here, we demonstrate that reducing klotho levels selectively in the CP disrupts the barrier between the immune system and the brain and promotes neuroinflammation.

Discussion Our findings suggest that klotho in the CP functions as a gatekeeper between the immune system and the CNS. The high level of klotho expressed in the CP of juvenile and adult mice, humans, and other mammals probably limits the penetration of this barrier by peripheral immune cells and suppresses the production of inflammatory mediators that could harm the CNS. We found that Klotho expression in the CP decreases with age and that experimental reduction of klotho levels specifically in this structure promotes the entry of peripheral macrophages and markedly increases the production of multiple proinflammatory mediators. Because similar proinflammatory changes occur during the natural aging process, the age-related decline in Klotho expression may contribute to the aging-related inflammation of the CNS. Because the CP produces and is bathed in CSF, inflammatory mediators produced in the CP are likely to reach other parts of the CNS via the CSF circulation (55), including the parenchyma of the brain and spinal cord, where they might contribute—by themselves or in combination with other factors—to the aging-related decline in neural functions (56). Indeed, experimental reduction of klotho in the CP exacerbated microglial activation in the hippocampus following peripheral injections of relatively low doses of LPS in the current study, suggesting that klotho depletion from the CP could contribute to the age-dependent “priming” of microglia for activation by peripheral infections (25, 38). Microglial activation and elevated IL-1β have been associated with deficits in synaptic plasticity in the hippocampus (57). A more detailed comparison of the results we obtained after experimentally reducing klotho levels in the CP of mice to those reported to spontaneously develop in the CP of aged mice reveals both similarities and differences. For example, klotho reduction in the CP increased the expression of IRF7, a key regulator of type I IFN (IFNα/β) responses, which have important roles in adaptive and innate immunity (58). IRF7 is also increased in the CP of aged mice and humans (32). In a similar vein, arginase-1 expression in the CP was increased in mice with selective klotho reduction in the CP (this study) as well as in aged mice (27). In contrast, we found increased levels of Icam1, Cxcl10, and Ccl17 mRNAs and of ICAM1 protein in the CP of mice with reduced klotho expression, whereas others found decreased levels of these gene products in the CP of aged mice (32). Some of these differences may reflect the fact that processes other than klotho reduction also affect the CP during aging and may override changes caused by klotho depletion. Other differences may have methodological or other reasons. For example, Baruch et al. reported increased IRF7 but decreased ICAM1 levels in the CP of aged mice (32), whereas CP levels of both proteins were increased in our aged mice, consistent with the results we obtained after reducing klotho levels in the CP of Klothoflox/flox mice by Cre injection. Our study provides insights into the mechanisms by which klotho may control the immune system/CNS interface in the CP. In combination, our in vivo and in vitro data support the hypothesis that klotho produced by CP epithelial cells suppresses the activation of the NLRP3 inflammasome in macrophages and possibly other cells, including the CP epithelial cells themselves, through FGF23/klotho signaling, counteracting 1,25-VD 3 , and inhibiting 1,25-VD 3 –dependent Txnip expression. TXNIP, which was increased in the CP after klotho reduction, can directly increase the expression of proinflammatory gene products that promote leukocyte infiltration into the CNS, including ICAM-1 (59). TXNIP also promotes activation of the NLRP3 inflammasome, a process that probably contributes causally to multiple aging-related deficits and aging-related diseases, including cognitive decline and neurodegenerative disorders (49, 60). Although TXNIP-dependent activation of the NLRP3 inflammasome is a plausible mechanism for many of the effects of klotho reduction we observed, our study was not designed to exclude alternative mechanisms by which klotho reduction in the CP might contribute to aging-related alterations of the CNS (5). Both klotho and the CP probably have diverse functions (3, 4, 6, 11, 33, 61). For example, klotho also attenuates the NF- κ B pathway (62), which could contribute to the increase in ICAM1 expression after klotho reduction. Similarly, increased expression of TXNIP could affect multiple processes besides inflammation (42, 45). Another limitation of our study is that we did not systematically compare the effects of klotho reduction in CPs of all ventricles, which share many but not all properties (6). In contrast to the genetic reduction of klotho in kl/kl mice, the injection of a Cre-expressing viral vector into Klothoflox/flox mice allowed us to (i) restrict klotho reduction specifically to the CP, (ii) avoid potential confounding effects of klotho reduction during early development, and (iii) lower klotho levels at distinct time points. Surprisingly, the viral vector approach revealed that klotho production by the CP is not needed to maintain normal klotho levels in the brain parenchyma, at least not in the hippocampus, where neurons produce low levels of klotho (18). Despite the distinct ways in which klotho levels were reduced in the two models, we confirmed in uninjected kl/kl mice several of the findings we obtained in Cre-injected Klothoflox/flox mice, highlighting the robustness of these findings. Alterations of the CP observed in both models included increased levels of Cyp27b1 mRNA, Txnip mRNA, and mRNAs encoding IFN-related factors, such as ICAM1, CCL17, and CXCR1. However, some changes we observed in Cre-injected Klothoflox/flox mice between 18 and 24 mo of age were not seen in uninjected kl/kl mice at 2 mo of age, including increased numbers of MAC-2- and Ly6C- positive cells and increased expression of the macrophage markers Nos2 and Arg1 in the CP. It is conceivable that these changes emerge over time, require aging-related cofactors that are missing in young kl/kl mice, or are prevented in kl/kl mice by the activation of compensatory processes during early development. Increased numbers of monocytes, macrophages, and T cells have been identified in the kidneys of heterozygous Klotho hypomorph (kl/+) mice (63). Our kl/kl mice had elevated TXNIP levels in the CP but not in the kidney. The latter discrepancy may reflect differences in cell type-specific inflammatory mechanisms, as reported for NLRP3 and receptors on macrophages and epithelial cells (64). Reduced klotho levels and increased IL-1β levels have also been observed in the CP of rats exposed to chronic unpredictable stress, an experimental paradigm that causes behavioral alterations in rodents reminiscent of endogenous depression in humans (19). IL-1β also elicits depression-related behaviors in rodents (65). Our findings may also be relevant to Alzheimer’s disease, in which klotho levels are decreased in the CSF (20, 23) and brain inflammation may have a critical pathogenic role (66⇓–68). Interestingly, global vitamin D 3 deficiency is a risk factor for age-related cognitive decline (69). Since vitamin D 3 has diverse effects throughout the body (69), it is tempting to speculate that high levels of klotho in the CP curtail immune invasion of the CNS via this important gateway by locally counteracting vitamin D 3 through FGF23/klotho signaling and reducing IFN/NLRP3-related gene expression, enabling the organism to benefit from useful vitamin D 3 effects elsewhere. Further investigation of klotho’s immune-regulatory roles in the CP may identify therapeutic strategies to block entry of harmful cells and factors into the CNS through the blood–CSF barrier, which might help counteract cognitive decline in elderly people and inflammaging-related neurological disorders.

Methods Mice and Treatments. Klothoflox/+ mice (24) were interbred to generate Klothoflox/flox and Klotho+/+ littermates on the same C57BL/6J background. CP-specific ablation of Klotho in Klothoflox/flox mice was achieved by stereotaxically injecting 2 μL of high-titer (1 to 10 × 1012/mL) AAV5-CMV-Cre-GFP (AAV5.CMV.HI.Cre.WPRE.SV40; Virovek) into the left lateral ventricle [coordinates: −1 mm mediolateral (M/L), −0.22 mm anteroposterior (A/P), and −2.5 mm dorsoventral (D/V) from the bregma]. Stereotaxic injections were carried out as described (70). WT mice injected with AAV5-CMV-Cre-GFP and in some cases Klothoflox/flox mice injected with AAV5-CMV-GFP (AAV5.CMV.HI.eGFP.WPRE.SV40; Virovek) served as negative controls. Overexpression of klotho in the hippocampus was achieved by injection of lentivirus encoding full-length klotho (Lenti-FUW2-klotho) into the dentate gyrus and CA1. The kl/kl mouse line (9) was obtained from M. Kuro-o (National Institute of Neuroscience, Kodaira, Tokyo) and the Nlrp3−/− line (51) from A. Ma (University of California, San Francisco, CA) Casp1/4−/− mice (52) were obtained from The Jackson Laboratory (strain 16621). Some mice received two separate i.p. injections of LPS (1 mg/kg; LPS-055: EB5; InvivoGen) 20 h apart and were perfused as described below 4 h after the second injection. Experimental and control groups were age-matched and littermates. Mice were maintained on a 12-h light/dark cycle, and experiments were conducted during the light cycle. Mice had free access to food (PicoLab Rodent Diet 20, 5053; LabDiet) and water, were housed 2 to 5 per cage. All procedures were approved by the Institutional Animal Care and Use Committee of the University of California, San Francisco. Isolation of Mouse Tissues. Mice were anesthetized with Avertin (0.025 mg/mL) and perfused transcardially with 0.9% saline. Hemibrains were removed and drop-fixed in 4% paraformaldehyde overnight at 4 °C or flash-frozen in liquid nitrogen and stored at −80 °C. Frozen hemibrains were thawed in cold 1× PBS containing Halt Protease and Phosphatase Inhibitor Mixture (78447; Thermo Fisher Scientific) and microdissected immediately. The intact choroid plexus (CP) was removed first to avoid contaminating other tissues. Microdissected tissues were immediately frozen on dry ice and kept at −80 °C. Primary Macrophage Cultures. Primary macrophages were prepared from 3- to 5-mo-old WT C57BL/6J mice as described (71). Femur bone marrow was extracted and mechanically dissociated in RPMI-1640 medium (Life Technologies). The cells were cultured in RPMI-1640, 10% FBS, 20 mM penicillin/streptomycin and 20 ng/mL macrophage colony-stimulating factor (ProSpec) at 2 × 106 cells per non-TC grade sterile 100-mm Petri dish (Corning). After 3 d, cells were dissociated in ice-cold PBS and replated in 12-well plates at 1 × 105 per well. Experiments were done on confluent cultures (typically days in vitro 7 to 8). Cultures were treated with ultrapure LPS (104 units/mL) for 16 to 24 h and harvested for analysis. Some cultures were incubated for an additional 4 h with ATP (1 mM; Sigma). In some cases, recombinant klotho (R&D Systems) and FGF-23 (R&D Systems) were added 1 h before LPS. After treatment, cells were washed once with PBS and collected for protein or mRNA analysis. Immunohistochemistry. Brain sections were prepared and immunostained as described (72⇓–74). Briefly, hemibrains were drop-fixed in 4% paraformaldehyde overnight, washed in cold PBS, equilibrated in 30% sucrose for ≥48 h, and stored at 4 °C. Hemibrains were sectioned (30 μm) on a freezing microtome, immunostained, and imaged with a digital microscope (BZ-9000; Keyence) or a laser-scanning microscope (LSM 880; Zeiss). Except for MHC II, antigen retrieval was performed using 10 mM citrate buffer at 105 °C for 20 min. After cooling at room temperature for at least 45 min, sections were incubated with 3% H 2 O 2 for 15 min to quench endogenous peroxidase, washed four times in PBS, incubated in blocking/permeabilization solution containing 10% normal goat serum and 0.3% Triton-X in PBS for ≥45 min, and incubated for ≥24 h with primary antibodies in 5% normal goat serum and 0.1% Triton X-100 in PBS at room temperature (dilutions provided in SI Appendix, Table S2). After three washes in PBS, sections were incubated with secondary antibodies (1:500; Invitrogen) and Hoechst 33342 (H33570; Thermo Scientific) diluted in 2% normal goat serum and 0.1% Triton X-100 in PBS for 2 h at room temperature. CLAUDIN-1 was detected with TSA-Plus cyanine 5 kits (PerkinElmer). Negative controls included omission of primary or secondary antibodies. High-resolution imaging was done with a Zeiss LSM 880 or a Keyence BZ-9000 automated microscope system. The Zeiss LSM 880 inverted scanning confocal microscope (Carl Zeiss Microscopy) was equipped with two photomultiplier tube (PMT) detectors, a high-sensitivity Gallium arsenide phosphide (GaAsP) detector, and a 32-GaAsP Airyscan superresolution detector and run by Zeiss Zen imaging software. The Keyence BZ-9000 inverted epifluorescence microscope was equipped with a 12-bit monochrome camera with red, green, and blue (RGB) capability. Unless indicated otherwise, all CP images were from CP in the lateral ventricles. For confocal microscopy, images were taken in z-stacks (2- to 4-μm steps) through immunoreactive areas. Z-stacks were analyzed with ImageJ or with Imaris software (Bitplane). For each section, numbers of cells positive for IRF7, ICAM1, MAC-2, or LY6C in the CP were normalized to the CP area in any given section; two to five sections were analyzed per mouse, and positive cells per area were averaged to generate a single value for each mouse (n). For quantitation of IBA-1 immunoreactivities, sections were stained as described above and imaged on a Versa Slide Scanner (Leica Systems) in z-stacks of 2-μm steps. The images were then batch-processed and analyzed using a macro developed in-house and the following functions. Areas of TMEM119 and IBA-1 immunoreactivity were measured by applying an adaptive threshold function (https://sites.google.com/site/qingzongtseng/adaptivethreshold). IBA-1–positive cells were counted automatically in the same images using the analyzed particle function of ImageJ. Western Blotting. CPs from the fourth and lateral ventricles were combined for each mouse (except for SI Appendix, Fig. S3 in which they were analyzed separately), homogenized in radioimmunoprecipitation assay (RIPA) buffer containing Halt Protease and Phosphatase Inhibitor Mixture (Thermo Scientific), and sonicated (Episonic 1000; Epigentek) at amplitude 40 for 5 min; protein concentrations were determined by the bicinchoninic acid (BCA) method. Equal amounts of total protein (10 to 20 µg for CP and 20 to 30 µg for hippocampus and kidney) were loaded in 1× NuPAGE LDS sample buffer (NP0007; Thermo Fisher Scientific) and 1× Sample Reducing Agent (NP0009; Thermo Fisher Scientific), separated on NuPAGE 4 to 12% Bis‐Tris gels (WT1403A; Thermo Fisher Scientific) at 200 V for 1.0 to 1.5 h in ice-cold 1× NuPAGE MOPS SDS running buffer (NP0001-02; Thermo Fisher Scientific). Gels were wet-transferred to nitrocellulose membranes (16 h, 25 V). The blots were then blocked with Odyssey Blocking Buffer (927-40000; LI-COR) for 1 h at room temperature, incubated with primary antibodies (SI Appendix, Table S2) overnight at 4 °C, washed with PBS containing 0.05% Tween 20 (PBST) three to four times for 5 min at room temperature, incubated with secondary antibodies conjugated to IRDye (0.1 µg/mL; Li-COR) for 1 h at room temperature, and washed in PBST three to four times for 5 min at room temperature. Protein bands were visualized with an Odyssey CLx Infrared Imaging System (LI-COR) and quantified with Image Studio Software (LI-COR) following the manufacturer’s instructions. The mean target protein:GAPDH ratio in WT mice was defined as 1.0. Quantitative Real-Time RT-PCR. Total RNA from mouse CPs from lateral and fourth ventricles or hippocampus was isolated with RNeasy Kits (74106; Qiagen) and reverse-transcribed with High-Capacity cDNA Reverse Transcription Kits (Thermo Fisher Scientific). After reverse transcription, quantitative real-time PCR was conducted with an ABI Prism 7900HT Sequence Detection System and SYBR Green Nucleic Acid Detection Kit or TaqMan Gene Expression Master Mix (Thermo Fisher Scientific) and TaqMan primers (Applied Biosystems) (SI Appendix, Table S3). Levels of each target mRNA were detected with FAM dye. Normalized relative target mRNA levels were calculated by the 2–∆∆C T method (75) using GAPDH mRNA levels (detected with VIC dye in the same well) as the internal reference and expressed relative to mean values in the control group. Primer sequences used to detect transcripts encoding distinct klotho isoforms with the SYBR Green PCR method are shown in SI Appendix, Table S4. Statistical Analysis. Prism (version 5; GraphPad), R (R Development Core Team, www.R-project.org/), and Microsoft Excel were used for statistical analyses. “n” refers to the number of mice for all figures except Fig. 6, in which “n” refers to independent experiments. Differences between genotypes and treatments were assessed by unpaired, two-tailed t test with Welch’s correction or by one-way or two-way ANOVA and Bonferroni, Holm-Sidak, or Tukey post hoc tests. P < 0.05 was considered significant. Results were collected blinded to genotype and treatment of mice and cell cultures. Biological units were randomized during assays, sampling, and analysis.

Acknowledgments We thank M. Kuro-O for kl/kl mice; A. Ma for Nlrp3−/− mice; S. Imai for discussion; X. Wang and X. Yu for technical support; S. Ordway for editorial review; and R. Mott for administrative assistance. This work was supported by funding (to L.M.) from the US National Institutes of Health Grant NS088532 and the Ray and Dagmar Dolby Family Fund. L.R.S. was supported by Ruth L. Kirschstein National Research Service Award NS093766.

Footnotes Author contributions: L. Zhu, L.R.S., and L.M. designed research; L. Zhu, L.R.S., D.K., K.H., and G.-Q.Y. performed research; L. Zhan and T.E.L. contributed new reagents/analytic tools; L. Zhu and L.R.S. analyzed data; and L. Zhu, L.R.S., and L.M. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

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