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Neurotoxic reactive astrocytes are induced by activated microglia

Nature volume 541, pages 481–487 (2017)Cite this article

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Abstract

Reactive astrocytes are strongly induced by central nervous system (CNS) injury and disease, but their role is poorly understood. Here we show that a subtype of reactive astrocytes, which we termed A1, is induced by classically activated neuroinflammatory microglia. We show that activated microglia induce A1 astrocytes by secreting Il-1α, TNF and C1q, and that these cytokines together are necessary and sufficient to induce A1 astrocytes. A1 astrocytes lose the ability to promote neuronal survival, outgrowth, synaptogenesis and phagocytosis, and induce the death of neurons and oligodendrocytes. Death of axotomized CNS neurons in vivo is prevented when the formation of A1 astrocytes is blocked. Finally, we show that A1 astrocytes are abundant in various human neurodegenerative diseases including Alzheimer’s, Huntington’s and Parkinson’s disease, amyotrophic lateral sclerosis and multiple sclerosis. Taken together these findings help to explain why CNS neurons die after axotomy, strongly suggest that A1 astrocytes contribute to the death of neurons and oligodendrocytes in neurodegenerative disorders, and provide opportunities for the development of new treatments for these diseases.

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Figure 1: A serum-free culture model for A1 astrocytes.
Figure 2: A1 astrocytes do not promote synapse formation or function.
Figure 3: A1 astrocytes lose phagocytic capacity.
Figure 4: An astrocyte-derived toxic factor promoting cell death.
Figure 5: A1 (C3-positive) reactive astrocytes in human disease.

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Acknowledgements

We thank R. Vance (UC Berkeley) for gifting Il-1a-/- mice and E. R. Stanley (Albert Einstein College of Medicine) for gifting Csf1r-/- mice. This work was supported by grants from the National Institutes of Health (R01 AG048814, B.A.B.; RO1 DA15043, B.A.B.; P50 NS38377, V.L.D. and T.M.D.) Christopher and Dana Reeve Foundation (B.A.B.), the Novartis Institute for Biomedical Research (B.A.B.), Dr. Miriam and Sheldon G. Adelson Medical Research Foundation (B.A.B.), the JPB Foundation (B.A.B., T.M.D.), the Cure Alzheimer’s Fund (B.A.B.), the Glenn Foundation (B.A.B.), the Esther B O’Keeffe Charitable Foundation (B.A.B.), the Maryland Stem Cell Research Fund (2013-MSCRFII-0105-00, V.L.D.; 2012-MSCRFII-0268-00, T.M.D.; 2013-MSCRFII-0105-00, T.M.D.; 2014-MSCRFF-0665, M.K.). S.A.L. was supported by a postdoctoral fellowship from the Australian National Health and Medical Research Council (GNT1052961), and the Glenn Foundation Glenn Award. L.E.C. was funded by a Merck Research Laboratories postdoctoral fellowship (administered by the Life Science Research Foundation). W.-S.C. was supported by a career transition grant from NEI (K99EY024690). C.J.B. was supported by a postdoctoral fellowship from Damon Runyon Cancer Research Foundation (DRG-2125-12). L.S. was supported by a postdoctoral fellowship from the German Research Foundation (DFG, SCHI 1330/1-1). T.M.D. is the Leonard and Madlyn Abramson Professor in Neurodegenerative Diseases. The authors (N.P., M.K., V.L.D. and T.M.D.) acknowledge the joint participation by the Adrienne Helis Malvin Medical Research Foundation through its direct engagement in the continuous active conduct of medical research in conjunction with The Johns Hopkins Hospital and the Johns Hopkins University School of Medicine and the Foundation’s Parkinson’s Disease Program M-2014. We thank the Stanford Alzheimer’s disease research Centre (AG047366), the Stanford Health Care Brain Bank, The Arizona Ageing & Disability Resource Centers (AG019610) and Banner Sun Health for providing control and AD brain samples. We thank R. Reynolds and D. Gveric for providing control and MS brain samples from the UK Multiple Sclerosis Tissue Bank, funded by the Multiple Sclerosis Society of Great Britain and Northern Ireland (registered charity 207495). We would like to thank O. Pletnikova and J. C. Troncoso from the Department of Pathology, Johns Hopkins University School of Medicine, for providing control and PD human sections. We thank the Neurological Foundation of New Zealand Human Brain Bank at the University of Auckland for Control and HD tissue sections for IHC analysis. We thank R. Myers at Boston University Medical Centre for control and HD tissue for q-RTPCR analysis. We thank J. Trojanoswki at the University of Pennsylvania Institute on Aging for AD and ALS tissue samples for in situ analysis. We thank A. Mosberger and A. Rosenthal for careful review of the manuscript. We thank T. Jessell and T. Maniatis for their insightful discussions on motor neuron subtypes. We thank V. and S. Coates for their generous support.

Author information

Authors and Affiliations

  1. Department of Neurobiology, Stanford University, School of Medicine, Stanford, 94305, California, USA

    Shane A. Liddelow, Kevin A. Guttenplan, Laura E. Clarke, Frederick C. Bennett, Mariko L. Bennett, Alexandra E. Münch & Ben A. Barres

  2. Department of Pharmacology and Therapeutics, University of Melbourne, Parkville, 3010, Victoria, Australia

    Shane A. Liddelow & Christopher J. Bohlen

  3. Department of Psychiatry and Behavioral Sciences, Stanford University, School of Medicine, Stanford, 94305, California, USA

    Frederick C. Bennett

  4. Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California San Francisco, San Francisco, 94143, California, USA

    Lucas Schirmer

  5. Department of Neurology, Klinikum rechts der Isar, Technical University of Munich, Munich, 81675, Germany

    Lucas Schirmer

  6. Department of Biological Sciences, Korea Advanced Institute of Science and Technology (KAIST), Daejeon, 34141, South Korea

    Won-Suk Chung

  7. Department of Neurology & Neurological Sciences, Stanford University, School of Medicine, Stanford, 94305, California, USA

    Todd C. Peterson & Marion S. Buckwalter

  8. Department of Neurology, F. M. Kirby Neurobiology Center, Boston Children’s Hospital, Boston, 02115, Massachusetts, USA

    Daniel K. Wilton, Arnaud Frouin & Beth Stevens

  9. Department of Microbiology and Immunology, Stanford University, School of Medicine, Stanford, 94305, California, USA

    Brooke A. Napier

  10. Neuroregeneration and Stem Cell Programs, Institute for Cell Engineering, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Nikhil Panicker, Manoj Kumar, Valina L. Dawson & Ted M. Dawson

  11. Department of Neurology, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Nikhil Panicker, Manoj Kumar, Valina L. Dawson & Ted M. Dawson

  12. Adrienne Helis Malvin Medical Research Foundation, New Orleans, Louisiana, 70130-2685, USA

    Nikhil Panicker, Manoj Kumar, Valina L. Dawson & Ted M. Dawson

  13. Departments of Pediatrics and Neurosurgery, University of California San Francisco, San Francisco, 94143, California, USA

    David H. Rowitch

  14. Department of Paediatrics, University of Cambridge, Cambridge, CB2 0AH, UK

    David H. Rowitch

  15. Department of Physiology, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Valina L. Dawson

  16. Solomon H. Snyder Department of Neuroscience, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Valina L. Dawson & Ted M. Dawson

  17. Department of Pharmacology and Molecular Sciences, Johns Hopkins University School of Medicine, Baltimore, 21205, Maryland, USA

    Ted M. Dawson

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  1. Shane A. Liddelow
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Contributions

S.A.L. and B.A.B. designed the experiments and wrote the paper. All authors reviewed and edited the manuscript. S.A.L. performed experiments and analysed data. K.G. performed proliferation assays and single-cell analysis. L.E.C. performed and analysed electrophysiology recordings and W.-S.C. performed and analysed in vivo astrocyte synapse pruning experiments. M.L.B. and S.A.L. performed optic nerve crushes. B.A.N. performed and analysed bacterial experiments. F.C.B. performed and analysed FACS experiments. T.C.P. performed stroke (MCAO) experiments. C.J.B. developed microglia culture systems. N.P. and M.K. differentiated hES cells to dopaminergic neurons for toxicity assays. Immunohistochemistry and analysis of human tissue was performed by L.S. (MS samples), S.A.L., D.K.W. and A.F. (AD), D.K.W. and A.F. (HD and ALS samples), and N.P. and M.K. (PD samples). qPCR analysis of human tissue was performed by S.A.L. (AD, HD, ALS and PD samples) and L.S. (MS samples). A.E.M. and K.G. provided technical support.

Corresponding author

Correspondence to Shane A. Liddelow.

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Competing interests

B.A.B. is a co-founder of Annexon Biosciences Inc., a company working to make new drugs for treatment of neurological diseases.

Additional information

Reviewer Information Nature thanks M. Freeman, S. Koizumi and the other anonymous reviewer(s) for their contribution to the peer review of this work.

Extended data figures and tables

Extended Data Figure 1 Csf1r−/− mice lack microglia and have no compensatory increase in brain myeloid cell populations after LPS or vehicle control injections.

a–c, Gating strategy (live, single cells) for subsequent analysis of surface protein immunostaining. d, e, Gating strategy for TMEM119+ (microglia) and CD45lowCD11b+ cells used for further analysis. f–h, Representative plots showing abundant macrophage populations in P8 wild-type mice: CD45lowTMEM119+/TMEM119−, and CD45high brain macrophages (f), CD11B+CD45low and CD11B+CD45high cells after saline (g) or LPS (h) injection. i–k, Representative plots showing near-complete absence of brain macrophages in Csf1r−/− mice: CD45lowTMEM119−TMEM119−, and CD45high brain macrophages (i), CD11B+CD45low and CD11B+CD45high cells after saline (j) and LPS (k) injection. l, Relative abundance of CD11B+CD45low macrophages after LPS or control injection in wild-type compared to Csf1r−/− mice, expressed as percentage of total gated events shown in a. m, Relative abundance of CD11B+CD45high cells after LPS treatment, normalized to saline control injection in wild-type and Csf1r−/− animals. n–p, Pexidartinib (PLX-3397)-treated adult mice have a pronounced reduction in the number of microglia and no increase in myeloid cell infiltration after LPS-treatment compared to vehicle control treatment. Representative plots showing abundant macrophage populations in P28 wild-type control mice: TMEM119+ microglia (n), CD11B+CD45low and CD11B+CD45high cells after saline (o) and LPS (p) injection. q–s, Representative plots showing a large reduction in macrophage populations after PLX-3397 treatment: TMEM119− microglia (q), CD11B+CD45low and CD11B+CD45high cells after saline (r) and LPS (s) injection. t, u, gating strategy for TMEM119+ (microglia) and CD45lowCD11b+ cells used for analysis. v, Relative abundance of CD11B+CD45low macrophages in wild-type compared to PLX-3397-treated mice, expressed as percentage of total gated events. w, Relative abundance of CD11B+CD45high cells after LPS treatment, normalized to saline control injection in wild-type and PLX-3397-treated animals. x, y, Fold change data from microfluidic qPCR analysis of wild-type and PLX-3397-treated mouse immunopanned astrocytes collected 24 h after i.p. injection with saline or LPS (5 mg kg−1). We found that we could only deplete approximately 95% of microglia using this drug (v), and it is probable that the remaining 5% that are sufficient to induce this strong level of A1 reactivity in astrocytes following LPS-induced neuroinflammation, also account for the death of retinal ganglion cells in optic nerve crush experiments (Figs 1e, 4l). n = 3–6 individual animals per treatment condition and genotype. Data are mean ± s.e.m. *P < 0.05, one-way ANOVA (l, v); P = 0.77 (m), P = 0.90 (w), Student’s t-test, compared to age-matched wild-type control.

Extended Data Figure 2 Screen for A1 mediators.

a, Immunopanning schematic for purification of astrocytes. These astrocytes retain their non-activated in vivo gene profiles. b, Purified cells were >99% pure with very little contamination from other central nervous system cells, as measured by qPCR for cell-type specific transcripts. c, Heat map of pan-reactive and A1- and A2-specific reactive transcript induction following treatment with a wide range of possible reactivity inducers. These factors did not produce an A1-astrocyte phenotype alone or in combination. n = 8 per experiment. *P < 0.05, one-way ANOVA (increase compared to non-reactive astrocytes).

Extended Data Figure 3 Screen for A1 mediators.

a, Fold change data from published microarray datasets of A1 (neuroinflammatory) reactive astrocytes. b–h, Microfluidic qPCR analysis of purified astrocytes treated with lipopolysaccharide (LPS)-activated microglia conditioned medium (b), non-activated microglia conditioned medium (c), a combination of Il-1α, TNF and C1q (d), LPS-activated microglia conditioned medium pre-treated with neutralizing antibodies to Il-1α, TNF and C1q (e), astrocytes treated with Il-1α, TNF and C1q and subsequently treated with FGF (f), microglia conditioned medium activated with interferon-γ (IFNγ, g), and with TNF (h). n = 6 per experiment. Data are mean ± s.e.m.

Extended Data Figure 4 Activation of microglia following systemic LPS injection in knockout mice.

a–c, Mice from global single knockouts of Il-1α (a), TNF (b), and C1q (c) were treated with LPS (5 mg/kg, i.p.) and microglia were collected 24 h later. Single-knockout animals still showed upregulation of many markers of microglial activation, as determined by qPCR. n = 3 for Il-1α and C1q, n = 5 for TNF. d, Quantitative PCR for microglia-derived A1-inducing molecules in the optic nerve of mice that received an optic nerve crush. Following crush, the optic nerve contained neuroinflammatory microglia, whereas injection of A1-astrocyte-neutralizing antibodies into the vitreous of the eye did not decrease microglial activation (however it did halt A1 astrocyte activation in the retina, see Fig. 4). Data are mean ± s.e.m. *P < 0.05, one-way ANOVA.

Extended Data Figure 5 A1 astrocytes are morphologically simple and do not promote synapse formation or neurite outgrowth.

a, In vivo immunofluorescence staining for the water channel AQP4 and GFAP. Saline-injected (control) mice showed robust AQP4 protein localization to astrocytic endfeet on blood vessels (white arrows), whereas LPS-injected mice showed a loss of polarization of AQP4 immunoreactivity, with bleeding of immunoreactivity away from endfeet (white arrows) and increased staining in other regions of the astrocyte (yellow arrowheads). Triple-knockout mice (Il1a−/−TNF−/−C1qa−/−) retained AQP4 immunoreactivity in endfeet following LPS-induced neuroinflammation (white arrows), although some low-level ectopic immunoreactivity was still seen (yellow arrowheads). b–e, Quantification of cell morphology of GFAP-stained cultured astrocytes in resting or A1 reactive state: cross-sectional area (b), number of primary processes extending from cell soma (c), number of terminal branchlets (d), ratio of terminal to primary processes (complexity score, e). f–h, Representative time-lapse tracing of control (f) and A1 (g) astrocytes. Quantification is shown in h. A1 astrocytes migrated approximately 75% less than control astrocytes over a 24-h period. n = 100 individual cells from at least 10 separate experiments. i, Total number of synapses normalized to each individual RGC. The number of synapses decreased after growth of RGCs with LPS-activated microglial conditioned medium (MCM)-activated A1 astrocyte conditioned medium (ACM), or Il-1α, TNF, C1q-activated (A1 astrocytes) was not different. j, Quantification of individual pre- and postsynaptic puncta. k, Total length of neurite growth from RGCs. l, Density of RGC processes in cultures used for the measurement of synapse number. There was no difference in neurite density close to RGC cell bodies (where synapse number measurements were made). n = 50 neurons in each treatment. m, Western blot analysis of proteoglycans secreted by control and A1 astrocytes. Conditioned medium from control astrocytes contained less chondroitin sulphate proteoglycans brevican, NG2, neurocan and versican, but contained higher levels of the heparan sulphate proteoglycans syndecan and glypican. *P < 0.05, Student’s t-test (d, k) or one-way ANOVA (all other panels). Scale bars, 100 μm. Data are mean ± s.e.m.

Extended Data Figure 6 P4 lateral geniculate nucleus astrocytes become A1 astrocytes following systemic LPS injection.

Fold change data from microfluidic qPCR analysis of astrocytes purified from dorsal lateral geniculate nucleus of P4 wild-type mice, 24 h after systemic injection with LPS (5 mg kg−1); n = 2.

Extended Data Figure 7 A1 astrocytes are strongly neurotoxic.

a, Quantification of dose-responsive cell death in retinal ganglion cells (RGCs) treated with astrocyte conditioned medium from cells treated with Il-1α, TNF or C1q alone, or combination of all three (A1 astrocyte conditioned medium, ACM) for 24 h. b, Death of RGCs was not due to a loss of trophic support, as treatment with 50% control ACM did not decrease viability. Similarly, treatment with a 50/50 mix of control and A1 ACM did not increase viability compared to A1-ACM-only treated cells. c, A1-ACM-induced RGC toxicity could be removed by heat inactivation or protease treatment. d, LPS treatment alone or microglia conditioned medium (MCM) treatment was unable to kill RGCs. ACM from astrocytes pre-treated with LPS, or non-activated MCM was also unable to kill RGCs. Only ACM from astrocytes treated with LPS-activated MCM or the combination of Il-1α, TNF and C1q (A1 astrocytes) were able to kill RGCs in culture. e–l, Cell viability of purified CNS cells treated with A1 ACM for 24 h: RGCs (e), hippocampal neurons (f), embryonic spinal motor neurons (g), oligodendrocyte precursor cells (OPCs, h), astrocytes (i), microglia/macrophages (j), endothelial cells (k) and pericytes (l). n = 4 for each experiment. m, Representative phase images showing the death of purified embryonic spinal motor neurons in culture over 48 h (ethidium homodimer stain in red shows DNA in dead cells). n, qPCR for motor neuronal subtype-specific transcripts after 120 h treatment with A1 ACM (50 μg ml−1). There was no decrease in levels of transcript for Nr2f2 (pre-ganglionic specific) and Wnt7a and Esrrg (gamma specific), suggesting these motor neuron subtypes are immune to A1-induced toxicity. n = 4 separate primary cultures. o, p, Representative images (o) and quantification (p) of terminal deoxynucleotidyl transferase (TdT) dUTP nick-end labelling (TUNEL) staining in the dentate gyrus for wild-type and Il1a−/−, TNF−/− or C1qa−/− individual knockout animals following systemic LPS injection. Individual knockout animals had far less TUNEL+ cells in the dentate gyrus (no cells in Il1a−/− or TNF−/− animals) than wild-type animals, suggesting A1-induced toxicity may be apoptosis (n = 5). q–s, Percentage growth rate of gram-negative bacterial cultures treated with A1 ACM for 16 h: B. thaliandensis (q), S. typhimurium (r), S. flexneri (s). n = 3; *P < 0.05, one-way ANOVA. Data are mean ± s.e.m.

Extended Data Figure 8 A1 astrocytes inhibit oligodendrocyte precursor cell proliferation, differentiation and migration.

a, Number of cells counted from phase-contrast images of oligodendrocyte precursor cells (OPCs) treated with control or A1-conditioned medium (ACM). b, EdU-ClickIt-assay-determined growth of OPCs treated with increasing concentrations of control and A1 ACM for 7 days. Both a and b show that A1 ACM decreases OPC proliferation compared to control. n = 6 separate primary cultures each. c–e, Representative images of time-lapse tracked OPC migration following treatment with control (c) and A1 (d) ACM, quantified in e. n = 100 cells from 10 separate experiments. f–h, qPCR shows no increase in the mature oligodendrocyte marker transcript Mbp in rat OPCs treated with A1 ACM, with no change in OPC marker Pdgfra and Cspg4 expression—evidence of a lack of differentiation into mature oligodendrocytes. Treatment of OPCs with control ACM did not delay their differentiation into mature oligodendrocytes (n = 2). i–k, Total number of terminal processes of rat oligodendrocyte lineage cells were counted as a measure of differentiation. Over 90% of cells differentiated by 24 h after removal of PDGFα when treated with control ACM (i). By contrast, treatment with a single dose (j) or daily doses (k) of A1 ACM delayed this level of differentiation up to 72 h following a single dose, or up to the limits of this experiment with chronic treatment. n = 6 separate experiments. l, Representative phase images and time scale for the oligodendrocyte differentiation assay (treated with control ACM). Scale bars, 100 μm (c, d) and 25 μm (l). *P < 0.05, one-way ANOVA, except for e (Student’s t-test). Data are mean ± s.e.m.

Extended Data Figure 9 Pharmacological blockade of an astrocyte-derived toxic factor.

a–j, Specific caspase inhibitory agents were tested whether these could block retinal ganglion cell (RGC) cell death. a, Pan-caspase inhibitor (Z-VAD-FMK). b, Caspase-1 inhibitor (Z-WEHD-FMK). c, Caspase-2 inhibitor (Z-VDVAD-FMK). d, Caspase-3 inhibitor (Z-DEVD-FMK). e, Caspase-4 inhibitor (Z-YVAD-FMK). f, Caspase-6 inhibitor (Z-VEID-FMK). g, Caspase-8 inhibitor (Z-IETD-FMK). h, Caspase-9 inhibitor (Z-LEHD-FMK). i, Caspase-10 inhibitor (Z-AEVD-FMK). j, Caspase-13 inhibitor (Z-LEED-FMK). Only caspase-2, caspase-3, casepase-4 and caspase-13 inhibition was able to minimize RGC toxicity induced by A1 ACM. Cleaved caspase-2 and -3 were detected in dying RGCs (Fig. 4). No cleaved caspase-4 or caspase-13 was detected in these cells. k, Necrostatin did not preserve RGC viability when cells were treated with A1 astrocyte conditioned medium (ACM). l–o, Glutamate excitotoxicity was checked by blocking AMPA receptors with the antagonist NBQX (l), NMDA antagonist D-AP5 (m) or kainite receptors with the antagonist UBP-296 (GluR5 selective, n) and UBP-302 (o)—all of which were ineffective. *P < 0.05, one-way ANOVA, n = 4 for each experiment. Data are mean ± s.e.m.

Extended Data Figure 10 Single-cell analysis of C3 expression in neuroinflammation and after ischaemic injury.

A1 astrocytes were induced with systemic injection of LPS (5 mg kg−1), and A2 astrocytes were induced with middle cerebral artery occlusion in Aldh1l1–eGFP mice. Individual astrocytes were collected via FACS and analysed with single-cell microfluidics for astrocyte reactive transcripts. a, Cassettes of pan-, A1-, and A2-specific gene transcripts used to determine polarization state of astrocyte reactivity. Upregulation of combinations of each of these cassettes of genes produces eight different possible gene profiles for astrocytes following injury. b, 24 h after LPS-induced systemic neuroinflammation, astrocytes were either non-reactive (no reactive genes upregulated), or fell into three forms of reactivity—all with A1 cassette genes upregulated. Numbers in parenthesis show the percentage of individual cells of each subtype expressed C3. c, 24 h after middle cerebral artery occlusion, both neuroinflammatory (A1 and A1-like) and ischaemic (A2 and A2-like) reactive cells were detected. No cells expressing A2 cassette transcripts were C3-positive, validating C3 as an appropriate marker for visualizing A1 astrocytes in disease. Segments of pie charts represent relative amounts of each subtype of astrocyte (control or reactive).

Extended Data Figure 11 Additional markers for reactive astrocytes in human post-mortem tissue samples.

a–h, Multiple Sclerosis (MS). a–c, Left, immunofluorescence staining showing CFB (a), MX1 (b), and C3 (c) co-localized with GFAP in cell bodies of reactive astrocytes in acute MS lesions (red arrows). Note the presence of A1-specific GFAP+ reactive astrocytes (C3+, CFB+, MX1+; red arrows) in close proximity to CD68+ phagocytes (activated microglia, macrophages; yellow arrowheads); A1-specific astrocytes are predominantly seen in high CD68+ density areas. C3−GFAP+ astrocytes (white arrows). Right, single channel and higher magnification images of selected areas of a–c. d, Immunohistochemical staining for C3 shows that it is strongly upregulated in astrocytes in active MS lesions. These astrocytes have a hypertrophic morphology with retracted processes (black arrows). Note hypercellularity indicating extensive infiltration by inflammatory cells in an active demyelinating MS lesion of subcortical white matter (compared to the luxol fast blue myelin stain of the lesion area in the right upper corner). e, C3 staining pattern in subcortical control white matter is mainly associated with blood vessels (stars). f, The A2-specific marker S100A10 did not co-localize with C3+ A1 astrocytes, although some S100A10+ cells were present in acute subcortical MS lesions. g, The number of C3+GFAP+ co-labelled cells was highest in acute active demyelinating lesions, however they were still present in chronic active and inactive lesions. h, There was a matching increase in C3 transcript in brains of patients with acute active demyelinating lesions compared to age-matched controls. FC, fold change. i, There was no difference in the expression of S100a10 in these same patient samples. n = 3–8 disease and 5–8 control for each experiment. Quantification was carried out on 5 fields of view and approximately 50 cells were surveyed per sample. j, k, Substantia nigra from patients with Parkinson’s disease (j) and age-matched controls (k). Additional representative images of C3+GFAP+ co-labelled cells in the substantia nigra of patients with Parkinson’s disease (j). No C3+ reactive astrocytes were seen in age-matched control substantia nigra (k). Scale bars, 100 μm (a–f), 20 μm (j, k and enlarged inserts). Data are mean ± s.e.m. *P < 0.05, one-way ANOVA, compared to age-matched control.

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Liddelow, S., Guttenplan, K., Clarke, L. et al. Neurotoxic reactive astrocytes are induced by activated microglia. Nature 541, 481–487 (2017). https://doi.org/10.1038/nature21029

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