Abstract
After injury, mammalian spinal cords develop scars to confine the lesion and prevent further damage. However, excessive scarring can hinder neural regeneration and functional recovery1,2. These competing actions underscore the importance of developing therapeutic strategies to dynamically modulate scar progression. Previous research on scarring has primarily focused on astrocytes, but recent evidence has suggested that ependymal cells also participate. Ependymal cells normally form the epithelial layer encasing the central canal, but they undergo massive proliferation and differentiation into astroglia following certain injuries, becoming a core scar component3,4,5,6,7. However, the mechanisms regulating ependymal proliferation in vivo remain unclear. Here we uncover an endogenous κ-opioid signalling pathway that controls ependymal proliferation. Specifically, we detect expression of the κ-opioid receptor, OPRK1, in a functionally under-characterized cell type known as cerebrospinal fluid-contacting neuron (CSF-cN). We also discover a neighbouring cell population that expresses the cognate ligand prodynorphin (PDYN). Whereas κ-opioids are typically considered inhibitory, they excite CSF-cNs to inhibit ependymal proliferation. Systemic administration of a κ-antagonist enhances ependymal proliferation in uninjured spinal cords in a CSF-cN-dependent manner. Moreover, a κ-agonist impairs ependymal proliferation, scar formation and motor function following injury. Together, our data suggest a paracrine signalling pathway in which PDYN+ cells tonically release κ-opioids to stimulate CSF-cNs and suppress ependymal proliferation, revealing an endogenous mechanism and potential pharmacological strategy for modulating scarring after spinal cord injury.
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Data availability
All data generated or analysed during this study are included in the manuscript and its extended data. The sequencing results have been deposited at the NCBI under the accession number GSE255883. The reference genome was built based on the annotated mouse reference genome (mm10) available as Genome assembly GRCm38 on the NCBI at: https://www.ncbi.nlm.nih.gov/datasets/genome/GCF_000001635.20/. Source data are provided with this paper.
References
Silver, J. & Miller, J. H. Regeneration beyond the glial scar. Nat. Rev. Neurosci. 5, 146–156 (2004).
Tran, A. P., Warren, P. M. & Silver, J. New insights into glial scar formation after spinal cord injury. Cell Tissue Res. 387, 319–336 (2021).
Johansson, C. B. et al. Identification of a neural stem cell in the adult mammalian central nervous system. Cell 96, 25–34 (1999).
Meletis, K. et al. Spinal cord injury reveals multilineage differentiation of ependymal cells. PLoS Biol. 6, e182 (2008).
Sabelström, H. et al. Resident neural stem cells restrict tissue damage and neuronal loss after spinal cord injury in mice. Science 342, 637–640 (2013).
Barnabé-Heider, F. et al. Origin of new glial cells in intact and injured adult spinal cord. Cell Stem Cell 7, 470–482 (2010).
Lacroix, S. et al. Central canal ependymal cells proliferate extensively in response to traumatic spinal cord injury but not demyelinating lesions. PLoS ONE 9, e85916 (2014).
New, L. E., Yanagawa, Y., McConkey, G. A., Deuchars, J. & Deuchars, S. A. GABAergic regulation of cell proliferation within the adult mouse spinal cord. Neuropharmacology 223, 109326 (2023).
Vigh, B., Vigh-Teichmann, I., Manzano e Silva, M. J. & van den Pol, A. N. Cerebrospinal fluid-contacting neurons of the central canal and terminal ventricle in various vertebrates. Cell Tissue Res. 231, 615–621 (1983).
Huang, A. L. et al. The cells and logic for mammalian sour taste detection. Nature 442, 934–938 (2006).
Orts-Del’immagine, A. et al. Properties of subependymal cerebrospinal fluid contacting neurones in the dorsal vagal complex of the mouse brainstem. J. Physiol. 590, 3719–3741 (2012).
Prendergast, A. E. et al. CSF-contacting neurons respond to Streptococcus pneumoniae and promote host survival during central nervous system infection. Curr. Biol. https://doi.org/10.1016/J.CUB.2023.01.039 (2023).
Böhm, U. L. et al. CSF-contacting neurons regulate locomotion by relaying mechanical stimuli to spinal circuits. Nat. Commun. 7, 10866 (2016).
Sternberg, J. R. et al. Pkd2l1 is required for mechanoception in cerebrospinal fluid-contacting neurons and maintenance of spine curvature. Nat. Commun. 9, 3804 (2018).
Orts-Del’Immagine, A. et al. A single polycystic kidney disease 2-like 1 channel opening acts as a spike generator in cerebrospinal fluid-contacting neurons of adult mouse brainstem. Neuropharmacology 101, 549–565 (2016).
Johnson, E. et al. Graded spikes differentially signal neurotransmitter input in cerebrospinal fluid contacting neurons of the mouse spinal cord. iScience 26, 105914 (2023).
Gerstmann, K. et al. The role of intraspinal sensory neurons in the control of quadrupedal locomotion. Curr. Biol. 32, 2442–2453.e4 (2022).
Djenoune, L. et al. The dual developmental origin of spinal cerebrospinal fluid-contacting neurons gives rise to distinct functional subtypes. Sci Rep. 7, 719 (2017).
Nakamura, Y. et al. Cerebrospinal fluid-contacting neuron tracing reveals structural and functional connectivity for locomotion in the mouse spinal cord. eLife 12, e83108 (2023).
Stoeckel, M.-E. et al. Cerebrospinal fluid-contacting neurons in the rat spinal cord, a γ-aminobutyric acidergic system expressing the P2X2 subunit of purinergic receptors, PSA-NCAM, and GAP-43 immunoreactivities: light and electron microscopic study. J. Comp. Neurol. 457, 159–174 (2003).
Chavkin, C. Dynorphin — still an extraordinarily potent opioid peptide. Mol. Pharmacol. 83, 729–736 (2013).
Khachaturian, H. et al. Dynorphin immunocytochemistry in the rat central nervous system. Peptides 3, 941–954 (1982).
Veldman, M. B. et al. Brainwide genetic sparse cell labeling to illuminate the morphology of neurons and glia with Cre-dependent MORF mice. Neuron 108, 111–127.e6 (2020).
Furube, E. et al. Neural stem cell phenotype of tanycyte-like ependymal cells in the circumventricular organs and central canal of adult mouse brain. Sci. Rep. 10, 2826 (2020).
Brust, T. F. Biased ligands at the κ opioid receptor: fine-tuning receptor pharmacology. Handb. Exp. Pharmacol. 271, 115–135 (2022).
Bruchas, M. R. & Chavkin, C. Kinase cascades and ligand-directed signaling at the κ opioid receptor. Psychopharmacology 210, 137–147 (2010).
Eriksson, P. S., Nilsson, M., W��gberg, M., Hansson, E. & Rönnbäck, L. κ-Opioid receptors on astrocytes stimulate l-type Ca2+ channels. Neuroscience 54, 401–407 (1993).
Gurwell, J. A. et al. κ-Opioid receptor expression defines a phenotypically distinct subpopulation of astroglia: relationship to Ca2+ mobilization, development, and the antiproliferative effect of opioids. Brain Res. 737, 175–187 (1996).
Pan, Z. Z. Opioid receptor-mediated enhancement of the hyperpolarization-activated current (Ih) through mobilization of intracellular calcium in rat nucleus raphe magnus. J. Physiol. 548, 765–775 (2003).
Lai, J. et al. Dynorphin A activates bradykinin receptors to maintain neuropathic pain. Nat. Neurosci. 9, 1534–1540 (2006).
Laughlin, T. M. et al. Spinally administered dynorphin A produces long-lasting allodynia: involvement of NMDA but not opioid receptors. Pain 72, 253–260 (1997).
Bakshi, R. & Faden, A. I. Competitive and non-competitive NMDA antagonists limit dynorphin A-induced rat hindlimb paralysis. Brain Res. 507, 1–5 (1990).
Zhang, S. et al. Dynorphin A as a potential endogenous ligand for four members of the opioid receptor gene family. J. Pharmacol. Exp. Ther. 286, 136–141 (1998).
Riondel, P. et al. Evidence for two subpopulations of cerebrospinal-fluid contacting neurons with opposite GABAergic signaling in adult mouse spinal cord. J. Neurosci. https://doi.org/10.1523/JNEUROSCI.2289-22.2024 (2024).
Corns, L. F. et al. Cholinergic enhancement of cell proliferation in the postnatal neurogenic niche of the mammalian spinal cord. Stem Cells 33, 2864–2876 (2015).
Hussein, S. A. Functional characterization of the TRP-type channel PKD2L1. ERA https://doi.org/10.7939/R3P84495F (2015).
Felix, R. Molecular regulation of voltage-gated Ca2+ channels. J. Recept. Signal Transduct. 25, 57–71 (2008).
Barber, R. P., Vaughn, J. E. & Roberts, E. The cytoarchitecture of GABAergic neurons in rat spinal cord. Brain Res. 238, 305–328 (1982).
Djenoune, L. et al. Investigation of spinal cerebrospinal fluid-contacting neurons expressing PKD2L1: evidence for a conserved system from fish to primates. Front. Neuroanat. 8, 26 (2014).
Matson, K. J. E. et al. Single cell atlas of spinal cord injury in mice reveals a pro-regenerative signature in spinocerebellar neurons. Nat. Commun. 13, 5628 (2022).
Ren, Y. et al. Ependymal cell contribution to scar formation after spinal cord injury is minimal, local and dependent on direct ependymal injury. Sci Rep. 7, 41122 (2017).
Corns, L. F., Deuchars, J. & Deuchars, S. A. GABAergic responses of mammalian ependymal cells in the central canal neurogenic niche of the postnatal spinal cord. Neurosci. Lett. 553, 57–62 (2013).
Kozono, H., Yoshitani, H. & Nakano, R. Post-marketing surveillance study of the safety and efficacy of nalfurafine hydrochloride (Remitch® capsules 2.5 μg) in 3,762 hemodialysis patients with intractable pruritus. Int. J. Nephrol. Renovasc. Dis. 11, 9–24 (2018).
Bloodgood, D. W. et al. κ Opioid receptor and dynorphin signaling in the central amygdala regulates alcohol intake. Mol. Psychiatry 26, 2187–2199 (2021).
Madisen, L. et al. A robust and high-throughput Cre reporting and characterization system for the whole mouse brain. Nat. Neurosci. 13, 133–140 (2010).
Krashes, M. J. et al. An excitatory paraventricular nucleus to AgRP neuron circuit that drives hunger. Nature 507, 238–242 (2014).
Gee, J. M. et al. Imaging activity in neurons and glia with a Polr2a-based and Cre-dependent GCaMP5G-IRES-tdTomato reporter mouse. Neuron 83, 1058–1072 (2014).
Buch, T. et al. A Cre-inducible diphtheria toxin receptor mediates cell lineage ablation after toxin administration. Nat. Methods 2, 419–426 (2005).
Arnold, K. et al. Sox2+ adult stem and progenitor cells are important for tissue regeneration and survival of mice. Cell Stem Cell 9, 317–329 (2011).
Edelstein, A. D. et al. Advanced methods of microscope control using μManager software. J. Biol. Methods 1, e10 (2014).
Renier, N. et al. iDISCO: a simple, rapid method to immunolabel large tissue samples for volume imaging. Cell 159, 896–910 (2014).
Acknowledgements
We thank C. Zuker for the Tg(PKD2L1-Cre) mouse line; R. Palmiter for the Pdynfl/fl mice; N. Ingolia for computational resources for sequencing analyses; K. Lindquist for statistical advice; J. Poblete for technical support; R. Nicoll, M. Beattie, J. Bresnahan, A. Basbaum, J. Braz, M. Bruchas, H. Ingraham, A. Alvarez-Buylla, M. Delling, N. Bellono, Z. Jiang, K. Yackle and all current members of the Julius laboratory for discussion and critical reading of the manuscript; and support from staff in UCSF’s core facilities, including the Laboratory for Cell Analysis (S. Elmes; NIH Cancer Center Support grant P30CA082103), the Center for Advanced Light Microscopy (D. Larsen, K. Herrington and S. Y. Kim; S10 Shared Instrumentation grant 1S10OD017993-01A1 for the Nikon CSU-W1 spinning disk confocal microscope), the Center for Advanced Technology (E. Chow and D. Martinez) as well as the Mouse Microsurgery Core (M. Looney and L. Qiu; financial support from the UCSF Bakar ImmunoX Initiative). This work was supported by a Howard Hughes Medical Institute Hanna Gray Fellowship and a Croucher Fellowship for Postdoctoral Research (to W.W.S.Y.), a Damon Runyon Cancer Research Foundation Fellowship (DRG-(2387-30) to K.K.T.), the UCSF Program for Breakthrough Biomedical Research: New Frontier Research Award (to D.J.) and NIH grants (R01EY030138 to X.D. and R35 NS105038 to D.J.).
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W.W.S.Y., K.K.T. and D.J. conceived and designed the experiments. W.W.S.Y. characterized the expression of the κ-opioid receptor and ligand, performed calcium imaging of CSF-cNs, and conducted the histological and functional analyses related to in vivo pharmacological interventions of the κ-opioid signalling pathway. W.W.S.Y. and K.K.T. performed the electrophysiological recordings of CSF-cNs. W.W.S.Y., K.K.T. and D.J. analysed the data. K.T. and X.D. contributed essential AAV resources. W.W.S.Y., K.K.T. and D.J. wrote the manuscript with input and suggestions from all authors.
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Extended data figures and tables
Extended Data Fig. 1 Characterization of CSF-cNs and PDYN+ cells in the spinal cord.
a, Labelling of spinal cord cells by the Tg(PKD2L1-Cre);Rosa26LSL-tdTomato mouse line. The tdTomato reporter (magenta) was expressed in many other spinal cord cells in addition to the CSF-cNs (magnified on right), which were identified by their strong immunostaining signal for PKD2L1 (green), particularly in their bulbous projections within the central canal (CC). b, Labelling of spinal cord cells by the Tg(PKD2L1-Cre) mouse line when tdTomato reporter was delivered by intracerebroventricular AAV injection after adulthood. The PHP.eB serotype labels cells beyond the ependymal region (e.g., arrow in magnified view on right). c, Projection pattern of CSF-cNs at different levels of the spinal cord. Coronal spinal cord and brain sections from a mouse that had received intracerebroventricular injection of an AAV carrying the PLAP transgene. Sections were stained with NBT/BCIP to reveal the ventral projections of CSF-cNs. d, Longitudinal projection of CSF-cNs. Sagittal section of the spinal cord showing the nerve bundle formed by CSF-cNs’ projections that travel rostrocaudally within the ventral white matter. Cell bodies of CSF-cNs are found in the ependyma. e, Morphology of a single PLAP-labelled CSF-cN. Spinal cord was cleared by the iDisco method51 after NBT/BCIP staining. f, Morphology of Pdyn+ cells. Arrowheads trace the long processes of sparsely labelled Pdyn+ cells situated at the dorsal or ventral pole of the ependyma. Dotted lines mark the boundary between grey and white matters. g, Schematic depicting the projection pattern of CSF-cNs (green) and PDYN+ cells (magenta). h, Spinal cord sections from PdynCre;Rosa26LSL-tdTomato mice immunostained with antibodies against known markers (cyan) for various subsets of ependymal cells. SOX2 is expressed in PDYN+ cells (magenta) in the ependymal region but not in PDYN+ cells in the dorsal horn. Ependymal PDYN+ cells are also positive for NESTIN but not GFAP. All experiments have been repeated for at least 3 times with similar results.
Extended Data Fig. 2 Expression of transcripts of interest in preparations of acutely dissociated and FACS-enriched CSF-cNs.
CSF-cNs were labelled by tdTomato via i.c.v. AAV injection and were fluorescently sorted for bulk mRNA sequencing. Columns represent 4 separate preparations with increasing level of enrichment in CSF-cNs, as reflected by the expression level of tdTomato transcript. Colour scale is based on median-of-ratios calculation by DESeq2. a, Marker genes for oligodendrocytes (Plp1 and Mbp), migroglia (P2ry12) and astrocytes (Gfap), showing the degree of glial contamination. b, Genes typically involved in GABA metabolism. c, Genes related to neurotransmission. d, Genes for receptors proposed to be sensitive to κ opioid ligands. e, TRP and Cav channel genes.
Extended Data Fig. 3 Ca2+ responses of GCaMP5G-expressing CSF-cNs to OPRK1 agonists.
ai, Example ΔF/F traces showing the responses of CSF-cNs to local application of the κ agonist, Nalfurafine, in the absence (black) or presence (red) of the antagonist, DIPPA, in the bath. Local application of a high K+ solution was used to reveal all responsive neurons. Each trace is from a single cell. aii, ΔF/F images for the spinal cord slices in ai. Images are temporal averages over 10 sec of baseline or for the duration of the stimuli. CC: central canal. Scale bars are 20 μm. aiii, Collective data comparing CSF-cNs’ responses to Nalfurafine in the absence (black) and presence (red) of DIPPA. Each dot shows the integral DYNA response of a single cell normalized to the high-K+ response. Two-sided Mann-Whitney test: P = 0.0445; n = 29 and 28 cells. b, Same as a except that BRL-52537 was used as the agonist and Nor-BNI as the antagonist. Two-sided Mann-Whitney test: P = 0.0150; n = 20 and 12 cells. In all bar graphs, data are mean ± s.d. *P ≤ 0.05.
Extended Data Fig. 4 Pharmacological experiments to delineate the downstream pathway of OPRK1 signaling.
a, CSF-cNs showed significant responses only to κ agonist (DYNA, 0.5 μM, 1 min) but not to agonists of the bradykinin receptors (bradykinin, 0.5 μM, 1 min), the delta opioid receptor (SNC162, 0.5 μM, 1 min) and the mu opioid receptor (Endomorphin-1, 0.5 μM, 1 min). Calcium imaging of acutely harvested spinal cord slices in the presence of TTX. Each colour represents one cell. Agonists were applied without gaps in the order displayed on the graph. Because comparisons were done within the same cell, responses were raw area under ΔF/F traces and were not baseline-subtracted nor normalized to high-K+ responses as in other figures. Repeated measures Friedman non-parametric test and Dunn’s posthoc pairwise comparisons with baseline: Bradykinin (P = 0.5466), SNC162 (P > 0.9999), Endomorphin-1 (P > 0.9999), and DYNA (P < 0.0001); n = 13. b, Normalized integral DYNA response (mean ± s.d.) as in Fig. 2e, but in the presence of various inhibitors or in different ionic conditions. Molecular targets of the drugs are indicated in brackets. Routes of drug application are detailed in Methods. Kruskal-Wallis non-parametric test and Dunn’s posthoc pairwise comparisons with control, which is same as –Nor-BNI in Fig. 2d and Fig. 2e: YM254890 (P < 0.0001), U73122 (P = 0.0133), U73343 (P = 0.3874, not significant), Thapsigargin (P > 0.9999, not significant), Ivabradine (P > 0.9999, not significant), Chelerythrine Cl (P = 0.0149), Nifedipine (P = 0.4256, not significant), ω-Agatoxin (P < 0.0001), ω-Conotoxin (P = 0.0005), SNX482 (P > 0.9999, not significant) and NNC 55-0396 (P = 0.0004). Numbers of cells analyzed are in brackets above bars. c, Proposed signalling pathway downstream of OPRK1 in CSF-cNs. *P ≤ 0.05; ***P ≤ 0.001; ****P ≤ 0.0001.
Extended Data Fig. 5 Voltage-clamp recording on CSF-cNs during DYNA application.
ai, Representative voltage-clamp recordings of CSF-cNs. Membrane potential was held at −80 mV in the absence or presence of the κ antagonist, Nor-BNI, in the bath. No macroscopic current was observed during local DYNA application (line above trace). aii, Expanded view of the boxed regions of traces in ai, showing single channel openings at baseline or during DYNA application. b, Example of a single channel opening event and a spontaneous postsynaptic event to show the clear distinction between the two waveforms. c, Amplitude histogram of spontaneous single channel opening events detected at baseline. Two peaks at amplitude ~5 pA and ~11 pA were detected. The ~11 pA events resembled those described in earlier reports11,15, which were shown to originate from PKD2L1 channels15. d, Open probability of the ~11 pA channel before and after DYNA application with or without Nor-BNI in bath. Each pair of light-coloured dots is from a single cell. Group averages are in dark colours. Repeated measures two-way ANOVA: DYNA × Nor-BNI (F1,10 = 0.01981, P = 0.8909, not significant); n = 6 and 5. We did not analyse the ~5 pA events because they were not clearly discernible from background noise. e, Rate of postsynaptic events before and during DYNA application in normal aCSF bath. Each pair of light-coloured dots is from a single cell. Group averages are in dark colours. Wilcoxon matched-pairs signed rank non-parametric test: P = 0.8750, not significant; n = 7 and 7.
Extended Data Fig. 6 Cav current recording from CSF-cNs during DYNA application.
a, Representative voltage-clamp recordings of CSF-cNs. Membrane potential was held at −100 mV, and 20 mV voltage steps from −100 mV to 60 mV were applied before, during, and after DYNA application. Voltage-gated sodium and potassium channels were inhibited by NMDG and TTX in the external solution and by caesium in the internal solution. Ba2+ (10 mM) was supplemented to the external buffer to increase the conductance of Cav channels. b, Current-voltage relationship recorded under the 3 conditions in a. The general Cav channel blocker, Cadmium (Cd2+), was added to one cell to verify that the recorded current was through Cav channels. Current responses were normalized to membrane capacitance, represented as mean ± s.d. Mixed-effect analyses with Šidák correction for pairwise comparisons: statistical significance reported in table; n = 10 for baseline and DYNA conditions and n = 8 for washout (data from 2 cells were excluded due to unstable recording). *P ≤ 0.05.
Extended Data Fig. 7 CSF-cNs, Pdyn+ cells and scar formation in injured spinal cords.
Following dorsal hemisection, no EdU signal (white) was detected in (a) CSF-cNs labelled in the Tg(PKD2L1-Cre) mouse line and (b) Pdyn+ cells labelled in the PdynCre line on Day 9. EdU injection scheme as in Fig. 4b. c, Immunohistochemical detection of PDYN in the ependymal region of sham-operated and injured mice 9 days after surgery. d, Effect of long-term κ agonist treatment (Nalfurafine, 0.027 mg/pellet with 60-day release rate, 5-6 weeks of treatment) on scar components. (Left) Representative images of the lesion sites in spinal cords harvested from placebo- or Nalfurafine-treated mice 5-8 weeks after dorsal hemisection. Sections were immunostained with antibody against GFAP (astrocytes), SOX9 (expressed in ependymal lineage), OLIG2 (oligodendrocytes) and CD68 (activated microglia and macrophages), which label components of scar tissues. (Right) Intensity of immunosignal or cell number was quantified and normalized to the volume of the region of interest. Two-sided Welch’s t-test: GFAP (P = 0.0005; n = 9 and 6), SOX9 (p = 0.0358; n = 12 and 10), and OLIG2 (P = 0.5756, not significant; n = 12 and 10). Two-sided Mann-Whitney test: CD68 (P = 0.9546, not significant; n = 9 and 6). *P ≤ 0.05; ***P ≤ 0.001.
Extended Data Fig. 8 Expression of the κ opioid receptor in various cell types.
a, Transcript expression of Oprk1 in various cell types that are known to participate in the injury response. Expression data and cluster identities were from ref. 40. The authors used contusion instead of dorsal hemisection as their injury model. The injury was inflicted at the thoracic level and spinal cord tissues were collected at the lumbar level. Color scale shows the average expression value. b, Protein expression of OPRK1 in various cell types identified by immunohistochemical markers. OPRK1 expression was detected only in CSF-cNs in the ependymal region (arrows) but not in other cell types in both sham-operated and injured animals 9 days after dorsal hemisection. Similar results were obtained from 3 experiments.
Extended Data Fig. 9 Effect of Nalfurafine (0.027 mg/pellet with 60-day release rate, 5-8 weeks of treatment) on rotarod performance.
Amount of time individual mice from each treatment group stayed on rotarod. Datapoints are the 3 highest scores; corresponding averages are represented by bars. The cutoff time is 10 min. Pie charts (black) indicate the fraction of animals that reached the cutoff time in at least 2 trials (i.e., censored). Number of animals tested are given in brackets.
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Yue, W.W.S., Touhara, K.K., Toma, K. et al. Endogenous opioid signalling regulates spinal ependymal cell proliferation. Nature (2024). https://doi.org/10.1038/s41586-024-07889-w
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DOI: https://doi.org/10.1038/s41586-024-07889-w
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