Epigenomic dynamics are associated with cell differentiation and maturation in the mammalian brain and have an essential role in regulating neuronal functions and animal behaviour5,6. Cytosine DNA methylation (5mC) is a stable covalent modification that persists in post-mitotic cells throughout their lifetime and is critical for proper gene regulation6. In mammalian genomes, 5mC occurs predominantly at CpG sites (mCG), showing dynamic patterns at regulatory elements with tissue and cell-type specificity1,6,7,8, modulating binding affinity of transcription factors9 and controlling gene transcription5. Non-CpG cytosines are also abundantly methylated (mCH, H denotes A, C, or T)—uniquely in neurons—in the mouse and human brain6,10, which can directly affect DNA binding of methyl CpG binding protein 2 (MeCP2)11,12,13, causing Rett syndrome14. Levels of mCH at gene bodies are anti-correlated with gene expression and show high heterogeneity across neuronal cell types1,7.

A deeper understanding of epigenomic diversity in the mouse brain provides a complementary approach to transcriptome-based profiling methods for identifying brain cell types and allows genome-wide prediction of the regulatory elements and transcriptional networks underlying this diversity. Previous studies have demonstrated the utility of studying brain cell types and regulatory diversity using single-nucleus methylome sequencing (snmC-seq)1. This study uses snmC-seq22 to perform thorough methylome profiling with detailed spatial dissection in the adult postnatal day 56 (P56) male mouse brain. In Li et al.3, the same tissue samples were profiled using single-nucleus assay for transposase-accessible chromatin using sequencing (snATAC-seq) to identify genome-wide accessible chromatin15, providing complementary epigenomic information to aid in cell-type-specific regulatory genome annotation. Moreover, to further study cis-regulatory elements and their potential target genes across the genome, we applied single-nucleus methylation and chromosome conformation capture sequencing (sn-m3C-seq)4 to profile the methylome and chromatin conformation in the same cells.

These epigenomic datasets provide a detailed and comprehensive census of the diversity of cell types across mouse brain regions, allowing identification of cell-type-specific regulatory elements and their candidate target genes and upstream transcription factors. Here we construct a single-cell base-resolution DNA methylation dataset containing 103,982 methylomes from 45 dissected brain regions and use an iterative analysis framework to identify 161 predicted mouse brain subtypes. Comparing subtype-level methylomes enables us to identify 3.9 million genomic regions showing cell-type-specific mCG variation, covering approximately 50% (1,240 Mb) of the mouse genome. We show that differentially methylated transcription factor genes and binding motifs can be associated with subtype taxonomy branches, allowing the prediction of cell-type gene regulatory programs specific for each developmental lineage. Integration of these data with cell clusters identified on the basis of chromatin accessibility validates most methylome-derived subtypes, enabling the prediction of 1.6 million enhancer-like genomic regions. We identify cis-regulatory interactions between enhancers and genes using computational prediction and single-cell chromatin conformation profiling (in the hippocampus (HIP)). We also identify spatial methylation gradients in cortical excitatory neurons and dentate gyrus granule cells and associated transcription factors and motifs. We apply an artificial neural network (ANN) model to precisely predict single-neuron cell-type identity and brain area spatial location using its methylome profile as input and develop the brain cell methylation viewer ( as a portal for querying and visualization of cell- and cluster-level methylation data.

Single-cell DNA methylome atlas

We used snmC-seq22 to profile genome-wide 5mC at single-cell resolution (Fig. 1a) across the cortex, HIP, striatum and pallidum (or cerebral nuclei, CNU), and olfactory areas (OLF) (Fig. 1b–e) using adult male C57BL/6 mice16. In total, we analysed 45 dissected regions in two replicates (Extended Data Fig. 1, Supplementary Table 2). Fluorescence-activated nuclei sorting (FANS) of antibody-labelled nuclei was applied to capture NeuN-positive neurons (NeuN+, 92% of neurons), while also sampling a smaller number of NeuN-negative (NeuN, 8% of neurons) non-neuronal cells (Fig. 1a). In total, we profiled the DNA methylomes of 103,982 single nuclei, yielding, on average, 1.5 million stringently filtered reads per cell (1.50 × 106 ± 0.58 × 106, mean ± s.d.) covering 6.2 ± 2.6% of the cytosines in the mouse genome in each cell. These enabled reliable quantification of the DNA methylation fraction for 25,905 ± 1,090 (95 ± 4%) 100-kb bins and 44,944 ± 4,438 (81 ± 8%) gene bodies (Extended Data Fig. 2a). The global methylation levels range from 0.2% to 7.6% in non-CpG sites and 61.6% to 88.8% in CpG sites (Extended Data Fig. 2b, c).

Fig. 1: A survey of single-cell DNA methylomes in the mouse brain.
figure 1

a, The workflow of dissection, FANS and snmC-seq2 sequencing. be, Dissected regions of isocortex (b), OLF (c), HIP (d) and CNU (e). f, Three-level UMAP from iterative analysis, colour coded as in be, panels show an example in which MSN-D1 neurons are separated into subtypes. g, Proportions of cells in clusters defined in the three-level iterative analysis. Brain atlas images in ad were created based on Wang et al.16 and © 2017 Allen Institute for Brain Science. Allen Brain Reference Atlas. Available from:

On the basis of the mCH and mCG profiles in 100-kb bins throughout the genome, we performed a three-level iterative clustering analysis to categorize the epigenomic cell populations (Fig. 1f, g). After quality control and preprocessing (Methods), in the first level (cell class), we clustered 103,982 cells as 67,472 (65%) excitatory neurons, 28,343 (27%) inhibitory neurons, and 8,167 (8%) non-neurons (Supplementary Table 3). The second round of iterative analysis of each cell class identified 41 cell major types in total (cluster size range 95–11,919), and the third round separated these major types further into 161 cell subtypes (cluster size range 12–6,551). All subtypes are highly conserved across replicates, and replicates from the same brain region are co-clustered compared with samples from other brain regions (Extended Data Fig. 2d–g).

The spatial distribution of each cell type is assessed based on where the cells were dissected (Supplementary Table 5). Here we used uniform manifold approximation and projection (UMAP)17 to visualize cell spatial locations (Fig. 1f, Extended Data Fig. 3) and major cell types (Extended Data Fig. 2h). Major non-neuronal cell types have a similar distribution across brain regions (Extended Data Fig. 1g), except adult neuron progenitors (ANPs). We found two subtypes of ANPs, presumably corresponding to neuronal precursors in the subgranular zone of the dentate gyrus (DG)18 (ANP anp-dg) and the rostral migratory stream18 in CNU and OLF (ANP anp-olf-cnu). Excitatory neurons from isocortex, OLF and HIP formed different major types, with some exceptions, potentially owing to overlaps in dissected regions (Supplementary Table 2). Cells from the isocortex were further separated on the basis of their projection types1,19,20. The intratelencephalic (IT) neurons from all cortical regions contain four major types corresponding to the laminar layers (L2/3, L4, L5 and L6), each of which includes cells from all cortical regions, except L4, which lack cells from the prefrontal cortex (PFC) and anterior cingulate area (ACA). Excitatory neurons from the HIP were further partitioned into major types corresponding to DG granule cells and different subfields of cornus ammonis (CA). We also identified major types from cortical subplate structures, including the claustrum (CLA) and endopiriform nucleus (EP) from isocortex and OLF dissections. GABAergic inhibitory neurons from isocortex and HIP cluster together into five major types, whereas interneurons from CNU and OLF group into nine major types.

In total, we identified 68 excitatory and 77 inhibitory subtypes (Fig. 2a, b, Supplementary Table 7). Although there is no one-to-one correspondence between subtypes and brain regions, individual subtypes show differential regional enrichment (Fig. 2a, b, top right) and distinct global mCH levels, ranging from 0.98% (DG dg-all) to 4.64% (PAL-Inh Chat, an inhibitory subtype in pallidum (PAL)) (Fig. 2a, b, bottom right). Specifically, isocortical excitatory subtypes usually consist of cells majorly derived from either the sensorimotor (primary motor (MOp), secondary motor (MOs), primary somatosensory (SSp), and secondary somatosensory (SSs) cortex), medial (PFC and ACA), or frontal areas (orbital (ORB) and agranular insular (AI) area). In the OLF, excitatory cells from the anterior olfactory nucleus (AON) and main olfactory bulb (MOB) are enriched in the subtype OLF-Exc Bmpr1b, whereas cells from the piriform area (PIR) are relatively enriched in the other OLF-Exc subtypes. Similarly, some inhibitory subtypes in CNU and OLF also correspond to different substructures in these two regions (Supplementary Note 1), indicating substantial spatial-related methylation diversity among CNU and OLF interneurons. By contrast, most caudal (CGE) or medial (MGE) ganglionic eminence-derived inhibitory subtypes contain cells derived predominantly from all cortical or hippocampal regions. To better demonstrate the unprecedented level of neuronal subtype and spatial diversity in their DNA methylomes, we provide a web application to interactively display this information at different granularity ( We also provide a detailed discussion of how exemplified subtypes correspond to cell types with known functional and spatial features (Extended Data Fig. 4) in Supplementary Note 1.

Fig. 2: Epigenomic diversity of neurons.
figure 2

a, b, Level 2 UMAP of excitatory (a) and inhibitory (b) neurons, coloured by subtype, dissection region and global mCH fraction. c, d, Integration UMAP of the HIP excitatory neurons profiled by snmC-seq2 (c) and snATAC-seq (d; shows pseudo-cells). e, Overlap score of a-types and m-types. f, Overlap of CG-DMR and ATAC peaks in matched subtypes. g, i, j, Integration t-SNE of ET-L5 neurons profiled by snmC-seq2 (g) and epi-retro-seq (i, j), coloured by dissection region. Three SSp- and MOp-enriched subtypes are labelled by their marker gene. j, Medulla projecting neurons from SSp or MOp only. h, Spatial composition of ET-L5 subtypes.

Consensus epigenomic profiles

Integrating single-cell datasets collected using different molecular profiling modalities can help to establish a consensus cell-type atlas20,21. By integrating the methylome data with the chromatin accessibility data profiled using snATAC-seq on the same brain samples from a parallel study3, the two modalities validated each other at the subtype level (Fig. 2c, d, Extended Data Fig. 5a–f, Supplementary Table 10). We then calculated overlap scores between the original methylation subtypes (m-types) and the chromatin accessibility subtypes (a-types), which further quantified the matching of subtypes between the two modalities (Fig. 2e, Extended Data Fig. 5e, Methods). Moreover, the mCG DMRs (see below) highly overlap with open chromatin peaks in the hippocampal subtypes (Fig. 2f). Their mCG fractions and chromatin accessibility levels show similar cell-type-specificity across hippocampal subtypes, confirming the correct match of cell-type identities (Extended Data Fig. 5f).

Projection specificity of ET-L5 neurons

To further infer the projection targets of cell subtypes, we integrated our extra-telencephalic (ET) L5 neurons with epi-retro-seq data22. Epi-retro-seq uses retrograde viral labelling to select neurons projecting to specific brain regions, followed by methylome analysis of their epigenetic subtypes. Cells from the same brain region of the two datasets are colocalized on t-distributed stochastic neighbour embedding (t-SNE) analysis, validating the subtypes’ spatial distribution (Fig. 2g–i, Extended Data Fig. 5g–i). The overlap scores between unbiased (snmC-seq2) and targeted (epi-retro-seq) profiling experiments (Extended Data Fig. 5j) indicate that some subtypes identified from the same cortical area show different projection specificity. For example, SSp and MOp neurons were mainly enriched in three subtypes marked by Kcnh1, Tmtc2 and Nectin1, respectively. However, neurons projecting to the medulla in the MOp and SSp only integrate with the subtype marked with Kcnh1 (Fig. 2j), suggesting that the subtypes identified in unbiased methylome profiling have distinct projection specificities.

Regulatory taxonomy of neuronal subtypes

Having developed a consensus map of cell types based on their DNA methylomes, we identified 16,451 differentially CH-methylated genes (CH-DMGs) and 3.9 million CG-differentially methylated regions (CG-DMRs, 624 ± 176 base pairs (bp) mean ± s.d.) between the subtypes (Extended Data Fig. 6, Methods, Supplementary Note 2). snmC-seq2 captures both cell-type-specific gene expression and predicted regulatory events1,2. Specifically, both gene body mCH and mCG negatively correlate with gene expression in neurons, with mCH showing a stronger correlation than mCG1,6,7,13. CG-DMRs provide predictions about cell-type-specific regulatory elements and transcription factors whose motifs enriched in these CG-DMRs predict the crucial regulators of the cell type1,7,8.

To further explore the gene regulatory relationship between neuronal subtypes, we constructed taxonomy trees for excitatory and inhibitory subtypes, based on gene body mCH of CH-DMGs (Extended Data Fig. 7a, b, Methods). The dendrogram structures represent the similarities between these discrete subtypes and may reflect the developmental history of neuronal type specification19,23. Next, we used both CH-DMGs and CG-DMRs to annotate the tree and explore the features specifying cell subtypes (excitatory in Fig. 3a–c and inhibitory in Extended Data Fig. 7c, d). Specifically, we calculated a branch-specific methylation impact score for each gene or transcription factor motif that summarizes all of the pairwise comparisons related to that branch (Extended Data Fig. 7e; Methods). The impact score ranges from 0 to 1, with a higher score predicting stronger functional relevance to the branch. We assign 6,038 unique genes to branches within the excitatory taxonomy (5,975 in inhibitory taxonomy), including 406 transcription factor genes (412 in inhibitory taxonomy) using genes with impact scores greater than 0.3. For example, motifs from the ROR (also known as NR1F) family were assigned to the branch that separates superficial layer IT neurons from deeper layer IT neurons (Fig. 3d–f, node 9), whereas motifs from the CUX family were assigned to the IT-L2/3 branch, separating it from IT-L4/5 neurons (Fig. 3d–f, node 11). Both of these families contain members, such as Cux1, Cux2 and Rorb, that show laminar expression in the corresponding layers and regulate cortical layer differentiation during development19.

Fig. 3: Relating genes and regulatory elements to cell subtype taxonomy.
figure 3

a, Schematic of the two characteristics contained in the methylome profiles. b, c, Pairwise CH-DMG (b) and CG-DMR (c) counts between 68 excitatory subtypes. c, In each CG-DMR set, we further identify differentially enriched motifs (left). TF, transcription factor. d, Excitatory subtype taxonomy tree. e, Top impact scores of ranked genes for the left and right branches of nodes 1–7 in d. The top four genes are transcription factor genes (bold); these are followed by other protein-coding genes. The scatter plots below show cells involved in each branch. f, Branch-specific transcription factor motif families. The zoomed UMAPs show individual transcription factor genes in those families, whose differential mCH fractions are concordant with their motif enrichment.

After impact score assignment, each branch of this taxonomy was associated with multiple transcription factor genes and motifs, which potentially function in combination to shape cell-type identities24 (Fig. 3e, f). For example, we focused on two brain structures of interest: the CLA and the EP25,26. At the major-cell-type level, distinct clusters are marked by Npsr1 (EP) and B3gat2 (CLA). The known EP and CLA marker transcription factor Nr4a225 also shows hypomethylation in both clusters compared to other clusters. Accordingly, the NR4A2 motif is also associated with a branch that splits CLA neurons from IT-L6 neurons (Fig. 3d–f, node 6). On another branch separating EP from CLA and IT-L6 neurons, genes for several transcription factors, including NF-1 family members Nfia and Nfib and the RFX family member Rfx3, together with corresponding motifs (Fig. 3d–f, node 5) rank near the top. Our findings suggest that these transcription factors may function together with Nr4a2, potentially separating EP neurons from CLA and IT-L6 neurons.

Beyond identifying specific cell-subtype characteristics, we derived total impact (TI) scores to summarize the methylation variation of genes and motifs to understand their relative importance in cell type diversification and function (Extended Data Fig. 7f–k, Supplementary Note 3). By comparing the TI scores of genes and motifs calculated from the inhibitory and excitatory taxonomies, we found that there were more transcription factor genes and motifs having large TI scores in both cell classes than in either one or the other (Extended Data Fig. 7f, i). For instance, Bcl11b distinguishes OLF-Exc and IT neurons in the excitatory lineage and distinguishes CGE-Lamp5 and CGE-Vip in the inhibitory lineage. Similarly, Satb1 separates IT–L4 from IT-L2/3 and MGE from CGE in excitatory and inhibitory cells. These findings indicate broad repurposing of transcription factors for cell-type specification among distinct developmental lineages.

Enhancer–gene Interactions

To systematically identify enhancer-like regions in specific cell types, we predicted enhancer-DMRs (eDMR) by integrating matched DNA methylome and chromatin accessibility profiles3 of 161 subtypes (Fig. 4a, Methods). We identified 1,612,198 eDMR (34% of CG-DMRs), 73% of which overlapped with separately identified snATAC-seq peaks (Fig. 4b). Fetal-enhancer DMRs (feDMR) (that is, eDMRs between development time points) of forebrain bulk tissues8 show high (88%) overlap with eDMRs. Surprisingly, the eDMRs also cover 74% of the feDMRs from other fetal tissues8, indicating extensive reuse of enhancer-like regulatory elements across mammalian tissue types (Fig. 4b).

Fig. 4: Gene–enhancer landscapes in neuronal subtypes.
figure 4

a, Schematic of enhancer calling using matched DNA methylome and chromatin accessibility subtype profiles. Corr., correlation. b, Overlap of regulatory elements identified in this study and other epigenomic studies (ATAC peaks3 and feDMRs8). c, DMR–DMG correlation and the distance between DMR centre and gene TSS; each point is a DMR–DMG pair coloured by kernel density. d, Percentage of positively correlated eDMRs that overlap with forebrain feDMRs in each gene. e, t-SNE of cells analysed by sn-m3C-seq coloured by assigned major cell types. f, Interaction level (z-score across rows and columns) of differential loops in eight clusters at 25-kb resolution. g, h, Epigenomic signatures surrounding Foxp1. g, Triangle heat maps showing CA1 and DG chromatin contacts and differential loops. h, Genome browser sections showing detailed mCG and ATAC profiles near anchors of four CA1-specific loops. Red rectangles indicate loop anchors and red arrows indicate notable regulatory elements.

Next, we examined the relationship between the cell-type-signature genes and their potential regulatory elements. We calculated the partial correlation between all DMG–DMR pairs within 1 Mb distance using methylation levels across 145 neuronal subtypes (Methods). We identified a total of 1,038,853 (64%) eDMRs that correlated with at least one gene (correlation >0.3 with empirical P <0.005, two-sided permutation test, Extended Data Fig. 8a). Notably, for those strongly positive-correlated DMR–DMG pairs (correlation >0.5), the DMRs are largely (63%) within 100 kb of the transcription start sites (TSSs) of the corresponding genes but are depleted from ±1 kb (Fig. 4c, Extended Data Fig. 8b), whereas for the negatively correlated DMR–DMG pairs, only 11% of DMRs are found within 100 kb of the TSS (Extended Data Fig. 8c).

Using the gene–enhancer interactions predicted by this correlation analysis, we assigned eDMRs to their target genes. The percentages of feDMR-overlapping eDMRs vary markedly among genes (Fig. 4d, Extended Data Fig. 8d, e). Of note, DMRs assigned to the same gene show different mCG specificity among subtypes. For example, Tle4-correlated eDMR could be partitioned into three groups (Extended Data Fig. 8e–g). One group (G2) of elements that displayed little diversity in bulk data showed highly specific mCG and open-chromatin signals in MSN–D1/D2 neurons, whereas another group (G3) was specific to CT–L6 neurons. These two groups of DMRs suggest that possible alternative regulatory elements are used to regulate the same gene in different cell types, although further experiments are required to validate this hypothesis.

Together, these analyses allow us to carefully chart the specificity of regulatory elements identified in bulk tissues to the subtype level. Besides, we identified many regulatory elements that show more restricted specificity (for example, eDMRs correlated with Tle4 in MSN-D1/D2), providing abundant candidates for further pursuing enhancer-driven adeno-associated viruses (AAVs) that target highly specific cell types27.

3D genome structure of hippocampus

Distal enhancers typically regulate gene expression through physical interaction with promoters28. Therefore, to examine whether physical chromatin contacts support our correlation-based predictions of enhancer–gene associations, we generated sn-m3C-seq4 data for 5,142 single nuclei from the HIP (152,000 contacts per cell on average). We assigned these cells, on the basis of the sn-m3C-seq data, to eight major cell types based on integration with the snmC-seq2 HIP data. In total, 19,151 chromosome loops were identified in at least one of the cell types at 25-kb resolution (range from 1,173 to 12,614 chromosome loops per cell type).

Using DG and CA1 as examples, a notably higher correlation was observed between enhancers and genes at loop anchors than between random enhancer–gene pairs (Extended Data Fig. 8h). Reciprocally, the enhancer–gene pairs showing stronger correlation with methylation were more likely to be found linked by chromosome loops or within the same looping region (Extended Data Fig. 8i). We also compared the concordance of methylation patterns between genes and enhancers linked by different methods and found the pairs linked by loop anchors or closest genes had the highest correlation of methylation (Extended Data Fig. 8j). Together, these analyses validate the physical proximity of enhancer–gene pairs predicted by our correlation-based method in specific cell types.

Additionally, we observed significant cell-type-specific 3D genome structures. The major cell types could be distinguished on UMAP embedding on the basis of chromosome interaction (Fig. 4e), indicating the dynamic nature of genome architecture across cell types. Among the 19,151 chromosome loops, 48.7% showed significantly different contact frequency between cell types (Fig. 4f). eDMRs were highly enriched at these differential loop anchors (Extended Data Fig. 8k). mCG levels at distal cis-elements are typically anti-correlated with enhancer activity8. Thus, we hypothesized that enhancers at differential loop anchors might also be hypomethylated in the corresponding cell type. Indeed, using the loops identified in DG and CA1 as examples, we observed that enhancers at the anchor of cell-type-specific loops show corresponding hypomethylation in the same cell type that the loop is specific to (Extended Data Fig. 8l).

Many differential loops were observed near marker genes of the corresponding cell type. For example, Foxp1, a gene for a CA1-specific transcription factor29, has chromosome loops surrounding its gene body in CA1 but not DG (Fig. 4g, h). eDMRs and open chromatin were observed at these loop anchors. Notably, three loops in CA1 anchored at the TSS of the same transcript of Foxp1 (Fig. 4h). Stronger demethylation and chromatin accessibility were also observed at the same transcript than in other transcripts (Fig. 4h, box E). These epigenetic patterns might suggest a specific transcript of Foxp1 (Foxp1-225) is selectively activated in CA1. by contrast, Lrrtm4, encoding a DG specific presynaptic protein that mediates excitatory synapse development30, shows extensive looping to distal elements in DG but not CA1 (Extended Data Fig. 8n). Notably, among 34 genes showing alternative loop usage, 20 genes expressed in both DG and CA131; for example, the TSS of Grm7 interacts with an upstream enhancer in DG and gene body enhancers in CA1 (Extended Data Fig. 8o).

mC gradients in IT neurons

Cortical excitatory IT neurons are classified into major types corresponding to their laminar layers: L2/3, L4, L5 and L6 (Fig. 5a). In agreement with the anti-correlation between transcript levels and DNA methylation, we found hypomethylation in IT neurons of the layer marker genes19 (Extended Data Fig. 9a). Furthermore, UMAP embedding (Fig. 5a) reveals a continuous gradient of IT neurons resembling the medial–lateral distribution of the cortical regions (Fig. 5b), strongly suggesting that the arealization information is well preserved in the DNA methylome.

Fig. 5: Brain-wide spatial gradients of DNA methylation.
figure 5

a, UMAP for cortex IT neurons coloured by dissection regions. b, The 21 cortical dissection regions organized by spatial axis. c, Normalized mCH fraction of spatial CH-DMGs, with the same layout as b. d, UMAPs for DG granule cells coloured by their cell global mCH fractions and dissection regions. A, anterior; P, posterior; D, dorsal; V, ventral. e, Compound figure showing four cells groups organized according to DG gradient and the two gradient DMR groups separated according to the sign of the correlation to the cell’s global mCH level. f, g, Bottom, expression (z-score across rows) of genes positively (f) or negatively (g) correlated with DMRs across cell types along the granule cell maturation pathway33. Top, genome browser views of representative genes.

To systematically explore the spatial gradient of DNA methylation, we merged the cells into spatial groups on the basis of their cortical layer and region and generated a taxonomy between them (Methods). The taxonomy split the cells into four layer groups, followed by cortical-region separation within each layer (Extended Data Fig. 9c), providing a clear structure for investigating layer-related or region-related methylation variation. Specifically, the layer-related transcription factors included many known laminar marker genes and their DNA-binding motifs (Extended Data Fig. 9d), whereas some also show regional specific methylation differences. For example, Cux1, encoding a homeobox transcription factor specific to L2/3 and L4 neurons, is hypomethylated in motor (MO) and somatosensory (SS) cortex, but is hypermethylated in L2/3 of other regions, in agreement with patterns from in situ hybridization32. Cux2, which encodes another homeobox transcription factor, does not show the same regional specificity (Extended Data Fig. 9a). We also identified genes for many additional transcription factors that showed cortical region specificity (Fig. 5c, Extended Data Fig. 9e). For example, Etv6 is only hypomethylated in medial dissection regions across layers, whereas Zic4 is hypermethylated in those regions. By contrast, Rora shows an anterior–posterior methylation gradient within the L4 and L5 cells. Together, these observed methylome spatial gradients demonstrated the value of our dataset for further exploring the cortical arealization with cell-type resolution.

mC gradients in DG granule cells

Global methylation gradients are observed within large cell types. For example, DG granule cells were continuously distributed in the UMAP embedding from low to high global mCH and mCG (Fig. 5d, global mCH fraction 0.5–1.9%, mCG fraction 69–79%). This gradient correlated with the anterior–posterior position of brain sections. Granule cells from the most posterior DG regions had higher global methylation than cells from anterior regions (Fig. 5d).

mCH accumulates throughout the genome during postnatal brain development6,8. We reasoned that DG granule cells, which are continuously replenished by ongoing neurogenesis throughout the lifespan, may accumulate mCH during their post-mitotic maturation. If so, global mCH should correlate with the age and maturity of granule cells. To investigate this, we divided DG granule cells into four groups on the basis of their global mCH levels and investigated regions of differential methylation between the groups. We identified 219,498 gradient CG-DMRs between the four groups, among which 139,387 showed a positive correlation with global mCH (+DMR), and 80,111 were negatively correlated (−DMR) (Fig. 5e). Notably, genes overlapping +DMRs or −DMRs have different annotated functions: genes enriched in +DMRs (+DMRgenes, n = 328) were associated with developmental processes, whereas those enriched in −DMRs (−DMRgenes, n = 112) were related to synaptic function (Extended Data Fig. 10a, b).

To further test the relationship between the +DMRgenes, −DMRgenes and DG development, we examined the expression patterns of these genes across time using a single-cell RNA-seq dataset that grouped DG cells into eight cell types, along their developmental trajectory from radial glia to mature granule cells33. The +DMRgenes were more highly expressed in immature cell types than in mature cell types (for example, Tcf4; Fig. 5f, Extended Data Fig. 10c), whereas the −DMRgenes showed the reverse trend (for example, Rfx3; Fig. 5g, Extended Data Fig. 10d). These results are consistent with the hypothesis that young DG granule cells have low global mCH and low methylation at genes associated with neural precursors. Conversely, older DG granule cells accumulate greater global mCH and have low methylation at genes associated with mature neurons. Notably, the global mCH levels also correlate with the brain dissections (Fig. 5d), indicating that the spatial axis can partially explain the methylation gradient (Supplementary Note 4).

Next, we investigated whether the global methylation level is correlated with 3D genome architecture. By plotting the chromatin interaction strength against the anchors’ genomic distance, we observed a higher proportion of short-range contacts and a smaller proportion of long-range contacts in the groups with higher global mCH (Extended Data Fig. 10f). Although compartment strengths were not correlated with the global methylation changes (Extended Data Fig. 10g), the number of intra-domain contacts was positively correlated with global mCH across single cells (Extended Data Fig. 10h). After normalizing for the effect of decay, we found that insulation scores at domain boundaries were significantly lower in the groups with high global mCH levels (Extended Data Fig. 10i; all P < 1 × 10−10, two-sided Wilcoxon signed-rank test). Together these suggest that local structures may be more condensed over flanking regions in the high-mCH cell groups.

Cell type and spatial prediction model

To further quantify the spatial and cell-type information encoded in a single cell’s DNA methylome, we built a multi-task deep ANN using cell-level methylome profiles from this study (Fig. 6a). Specifically, mCH levels of 100-kb bins were used to train and test the network with fivefold cross-validation (Method). The ANN predicted neuronal subtype identity and spatial location simultaneously for each testing cell with 95% and 89% accuracy, respectively (Fig. 6b–d). Notably, the location prediction accuracy of the ANN was higher than using only the spatial distribution information of subtypes (overall increased by 38%, Extended Data Fig. 11c), suggesting that spatial diversity is well-preserved in the neuronal DNA methylome. We also notice higher levels of errors in location prediction of some cell types, especially in the cortical MGE and CGE inhibitory neurons (Fig. 6c, Extended Data Fig. 11c). This finding is consistent with previous transcriptome-based studies19,31, suggesting these neurons do not display strong cortical region specificity. Many cell-type marker genes are also enriched in features that capture most spatial information (Fig. 6e, f). For example, besides distinguishing CT-L6 neurons from other cell types, Foxp2 shows notable mCH differences among dissected regions within CT-L6 (Fig. 6f). Notably, we also observed the moderate spatial specificity of astrocytes and oligodendrocytes using a separate model trained with methylomes of non-neuronal cells (Supplementary Note 5).

Fig. 6: A methylome-based predictive model captures both cellular and spatial characteristics of neurons.
figure 6

a, Schematic of the model that predicts both cell-type identity and spatial origin. b, c, Model performance on the prediction of cell subtypes (b), and dissection regions (c). Per cent accuracy is shown. d, Examples of using the model to predict cell spatial origin (maximum prediction probability in parentheses). e, Evaluation of importance of features (principal components) for spatial origin prediction accuracy. ‘Acc. Δ%’ denotes the average prediction accuracy decrease percentage. f, The Foxp2 gene body mCH fraction in each cortical dissection region group.


In this Article, we present a single-cell DNA methylomic atlas of the mouse brain with detailed spatial dissection. This comprehensive dataset enables high-throughput cell-type classification, marker gene prediction and identification of regulatory elements. The three-level iterative clustering defined 161 subtypes representing excitatory (68), inhibitory (77) and non-neuronal cells (16). The development of a hierarchical taxonomic architecture for cell subtypes on the basis of CH-DMGs allowed us to assign specific genes and transcription factor binding motifs to taxonomy branches using the methylation impact score. These assignments describe cell-type specificity at different levels, potentially relating to different developmental stages of each neuronal lineage. Notably, we found that transcription factor genes and their corresponding DNA-binding motifs were co-associated with the same branch in the taxonomy, providing a rich source of candidate transcription factors for future study.

Through integration with snATAC-seq3, we matched subtypes classified in both epigenomic modalities and used the combined information to predict 1.6 million active-enhancer-like eDMRs, including 72% of cell-type-specific elements missed from previous tissue-level bulk studies8. To examine the associations of eDMRs and their targeting genes, we applied multi-omic methods to establish an eDMR–gene landscape using correlation-based prediction and chromatin conformation profiling using sn-m3C-seq, resulting in the identification of chromatin loops between eDMRs and their potential targeting genes in specific cell types.

Our brain-wide epigenomic dataset reveals extraordinary spatial diversity encoded in the DNA methylomes of neurons. The ANN trained on the single-cell methylome profiles accurately reproduced the detailed brain-dissection information within most subtypes, indicating the existence of large spatial methylation gradients throughout the brain. Echoing cortex development studies34, glutamatergic neurons are regionalized by a protomap formed from an early developmental gradient of transcription factor expression. Similarly, we observed that many transcription factor genes and their corresponding DNA-binding motifs showed gradients of DNA methylation in adult IT neurons from distinct cortical regions. Additionally, we also found intra-subtype methylation gradients in DG granule cells that correlate with the spatial axis in the DG. These gradient-related CG-DMRs are enriched in essential neurodevelopmental and synaptic genes33,35, suggesting that these spatially resolved DNA methylation gradients reflect past regulatory events occurring during brain maturation. We qualify our findings by noting that snmC-seq2 is a sodium bisulfite-based method and cannot distinguish between 5-methylcytosine and 5-hydroxymethylcytosine, which has been shown to accumulate in some brain regions36. New methods will be needed to simultaneously measure the full complement of cytosine base modifications at the single-cell level.

Overall, our analysis highlights the power of this dataset power for characterizing cell types using gene activity information from both coding regions and the regulatory elements in the non-coding regions of the genome. This comprehensive epigenomic dataset provides a valuable resource for answering fundamental questions about gene regulation in specifying cell-type spatial diversity and provides the raw material to develop new genetic tools for targeting specific cell types and functional testing.


Mouse brain tissues

All experimental procedures using live animals were approved by the Salk Institute Animal Care and Use Committee under protocol number 18-00006. Adult (P56) C57BL/6J male mice were purchased from Jackson Laboratories and maintained in the Salk animal barrier facility on 12 h dark-light cycles with food ad libitum for a maximum of 10 days. Brains were extracted and sliced coronally at 600 μm from the frontal pole across the whole brain (for a total of 18 slices) in an ice-cold dissection buffer containing 2.5 mM KCl, 0.5 mM CaCl2, 7 mM MgCl2, 1.25 mM NaH2PO4, 110 mM sucrose, 10 mM glucose and 25 mM NaHCO3. The solution was kept ice-cold and bubbled with 95% O2, 5% CO2 for at least 15 min before starting the slicing procedure. Slices were kept in 12-well plates containing ice-cold dissection buffers (for a maximum of 20 min) until dissection aided by an SZX16 Olympus microscope equipped with an SDF PLAPO 1XPF objective. Olympus cellSens Dimension 1.8 was used for image acquisition. Each brain region was dissected from slices along the anterior-posterior axis according to the Allen Brain reference Atlas CCFv316 (see Extended Data Fig. 1 for the depiction of a posterior view of each coronal slice). Slices were kept in ice-cold dissection media during dissection and immediately frozen in dry ice for posterior pooling and nuclei production. For nuclei isolation, each dissected region was pooled from 6–30 animals, and two biological replicas were processed for each slice.

Fluorescence-activated nuclei sorting

Nuclei were isolated as previously described1,6. Isolated nuclei were labelled by incubation with 1:1,000 dilution of Alexa Fluor 488-conjugated anti-NeuN antibody (MAB377X, Millipore) and a 1:1,000 dilution of Hoechst 33342 at 4 °C for 1 h with continuous shaking. FANS of single nuclei was performed using a BD Influx sorter with an 85-μm nozzle at 22.5 PSI sheath pressure. Single nuclei were sorted into each well of a 384-well plate preloaded with 2 μl of proteinase K digestion buffer (1 μl M-Digestion Buffer (Zymo, D5021-9), 0.1 μl of 20 μg μl−1 proteinase K and 0.9 μl H2O). The alignment of the receiving 384-well plate was performed by sorting sheath flow into wells of an empty plate and making adjustments based on the liquid drop position. Single-cell (one-drop single) mode was selected to ensure the stringency of sorting. For each 384-well plate, columns 1–22 were sorted with NeuN+ (488+) gate, and column 23-24 with NeuN (488−) gate, reaching an 11:1 ratio of NeuN+ to NeuN nuclei. BD Influx Software v1.2.0.142 was used to select cell populations.

Library preparation and Illumina sequencing

Detailed methods for bisulfite conversion and library preparation were previously described for snmC-seq21,2. The snmC-seq2 and sn-m3C-seq (see below) libraries generated from mouse brain tissues were sequenced using an Illumina Novaseq 6000 instrument with S4 flow cells using the 150-bp paired-end mode. Freedom EVOware v2.7 was used for library preparation, and Illumina MiSeq control software v3.1.0.13 and NovaSeq 6000 control software v1.6.0/Real-Time Analysis (RTA) v3.4.4 were used for sequencing.

The sn-m3C-seq specific steps of library preparation

Single-nucleus methyl-3C sequencing (sn-m3C-seq) was performed as previously described4. In brief, the same batch of dissected tissue samples from the dorsal dentate gyrus (DG-1 and DG-2, Supplementary Table 2), ventral dentate gyrus (DG-3 and DG-4), dorsal HIP (CA-1 and CA-2), and ventral HIP (CA-3 and CA-4), were frozen in liquid nitrogen. The samples were then pulverized while frozen using a mortar and pestle, and then immediately fixed with 2% formaldehyde in DPBS for 10 min. The samples were quenched with 0.2 M glycine and stored at −80 °C until ready for further processing. After isolating nuclei as previously described4, nuclei were digested overnight with NlaIII and ligated for 4 h. Nuclei were then stained with Hoechst 33342 (but not stained with NeuN antibody) and filtered through a 0.2-μm filter, and sorted similarly to the snmC-seq2 samples. Libraries were generated using the snmC-seq2 method.

Mouse brain region nomenclature

The mouse brain dissection and naming of anatomical structures in this study followed the Allen Mouse Brain common coordinate framework (CCF)16. On the basis of the hierarchical structure of the Allen CCF, we used a three-level spatial region organization to facilitate description: (1) the major region, for example, isocortex, HIP; (2) the sub-region, for example, MOp, SSp, within isocortex; (3) the dissection region, for example, MOp-1 and MOp-2, within MOp. Supplementary Table 1 contains the full names of all abbreviations used in this study. All brain atlas images were created based on Wang et al.16 and ©2017 Allen Institute for Brain Science. Allen Brain Reference Atlas. Available from:

Analysis stages

The following method sections were divided into three stages. The first stage, ‘Mapping and feature generation’, describes mapping and generating files in the single-cell methylation-specific data format. The second stage, ‘Clustering related’, describes clustering, identifying DMGs, or integrating other datasets, which all happened at the single-cell level. The third stage, ‘Cell-type-specific regulatory elements’, describes the identification of putative cell-type-specific regulatory elements using cluster-merged methylomes. Other figure-specific analysis topics may combine results from more than one stage.

Mapping and feature generation

Mapping and feature-count pipeline

We implemented a versatile mapping pipeline, YAP (, for all the single-cell-methylome-based technologies developed by our group1,2,37. The main steps of this pipeline include: (1) demultiplexing FASTQ files into single cells; (2) reads level quality control (QC); (3) mapping; (4) BAM file processing and QC; and (5) final molecular profile generation. The details of the five steps for snmC-seq2 were previously described2. We mapped all of the reads to the mouse mm10 genome. We calculated the methylcytosine counts and total cytosine counts for two sets of genomic regions in each cell after mapping. Non-overlapping chromosome 100-kb bins of the mm10 genome (generated by “bedtools makewindows -w 100000”) were used for clustering analysis and ANN model training, and the gene body regions ±2 kb defined by the mouse GENCODE vm22 were used for cluster annotation and integration with other modalities.

sn-m3C-seq-specific steps or read mapping and chromatin contact analysis

Methylome sequencing reads were mapped following the TAURUS-MH pipeline, as previously described4. Specifically, reads were trimmed for Illumina adaptors, and then an additional 10 bp was trimmed on both sides. Then R1 and R2 reads were mapped separately to the mm10 genome using Bismark with Bowtie. The unmapped reads were collected and split into shorter reads representing the first 40 bp, the last 40 bp, and the middle part of the original reads (if read length >80 bp after trimming). The split reads were mapped again using Bismark with Bowtie. The reads with MAPQ <10 were removed. The filtered bam files from split and unsplit R1 and R2 reads were deduplicated with Picard and merged into a single bam file to generate the methylation data. Methylpy (v1.4.2)38 was used to generate an ALLC file (base-level methylation counts) from the bam file for every single cell. We paired the R1 and R2 bam files where each read-pair represents a potential contact to generate the Hi-C contact map. For generating contact files, read pairs where the two ends mapped within 1 kbp of each other, were removed.

Clustering-related methods

Single-cell methylome data quality control and preprocessing

Cell filtering. We filtered the cells on the basis of these main mapping metrics: (1) mCCC level <0.03; (2) overall mCG level >0.5; (3) overall mCH level <0.2; (4) total final reads >500,000; and (5) Bismark mapping rate >0.5. Other metrics such as genome coverage, PCR duplicates rate and index ratio were also generated and evaluated during filtering. However, after removing outliers with the main metrics 1–5, few additional outliers were found. Note the mCCC level is used as the estimation of the upper bound of bisulfite non-conversion rate1.

Feature filtering. 100 kb genomic bin features were filtered by removing bins with mean total cytosine base calls <250 (low coverage) or >3,000 (unusually high-coverage regions). Regions that overlap with the ENCODE blacklist39 were also excluded from further analysis.

Computation and normalization of the methylation level. For CG and CH methylation, the methylation level computation from the methylcytosine and total cytosine matrices contains two steps: (1) prior estimation for the beta-binomial distribution, and (2) posterior level calculation and normalization per cell.

Step 1: for each cell, we calculated the sample mean \(m\) and variance \(v\) of the raw methylcytosine level (mc/cov), where cov is the total cytosine base coverage and mc is the methylcytosine base coverage, for each sequence context (CG or CH). The shape parameters (α, β) of the beta distribution were then estimated using the method of moments:

$$\alpha =m(m(1-m)/v-1)$$
$$\beta =(1-m)(m(1-m)/v-1)$$

This approach used different priors for different methylation types for each cell and used weaker priors to cells with more information (higher raw variance).

Step 2: we then calculated the posterior: \(\widehat{{\rm{mc}}}=\frac{\alpha +{\rm{mc}}\,}{\alpha +\beta +{\rm{cov}}}\) for all bins in each cell. Like the counts per million reads (CPM) normalization in the single-cell RNA-seq analysis, we normalized this posterior methylation ratio by the cell’s global mean methylation, m = α/(α + β). Thus, all the posterior \(\widehat{{\rm{mc}}}\) values with 0 cov will have a constant value of 1 after normalization. The resulting normalized mc level matrix contains no NA (not available) value, and features with lower cov tend to have a mean value close to 1.

Selection of highly variable features. Highly variable methylation features were selected with a modified approach using the scanpy.pp.highly_variable_genes function from the scanpy 1.4.3 package40. In brief, the scanpy.pp.highly_variable_genes function normalized the dispersion of a gene by scaling with the mean and standard deviation of the dispersions for genes falling into a given bin for mean expression of genes. In our modified approach, we reasoned that both the mean methylation level and the mean cov of a feature (100 kb bin or gene) could impact mc level dispersion. We grouped features that fall into a combined bin of mean and cov. We then normalized the dispersion within each mean–cov group. After dispersion normalization, we selected the top 3,000 features based on normalized dispersion for clustering analysis.

Dimension reduction and combination of different mC types. For each selected feature, mc levels were scaled to unit variance and zero mean. We then performed principal component analysis (PCA) on the scaled mc level matrix. The number of principal components (PCs) was selected by inspecting the variance ratio of each PC using the elbow method. The CH and CG PCs were then concatenated together for further analysis in clustering and manifold learning (Supplementary Table 6 for parameters of PCA and clustering analysis).

Consensus clustering

Consensus clustering on concatenated PCs. We used a consensus clustering approach based on multiple Leiden clustering41 over k-nearest neighbour (KNN) graph to account for the randomness of the Leiden clustering algorithms. After selecting dominant PCs from PCA in both mCH and mCG matrices, we concatenated the PCs together to construct a KNN graph using scanpy.pp.neighbours with Euclidean distance. Given fixed resolution parameters, we repeated the Leiden clustering 300 times on the KNN graph with different random starts and combined these cluster assignments as a new feature matrix, where each single Leiden result is a feature. We then used the outlier-aware DBSCAN algorithm from the scikit-learn package to perform consensus clustering over the Leiden feature matrix using the hamming distance. Different epsilon parameters of DBSCAN are traversed to generate consensus cluster versions with the number of clusters that range from the minimum to the maximum number of clusters observed in the multiple Leiden runs. Each version contained a few outliers; these usually fall into three categories: (1) cells located between two clusters had gradient differences instead of clear borders, for example, border of IT layers; (2) cells with a low number of reads potentially lack information in essential features to determine the specific cluster; and (3) cells with a high number of reads that were potential doublets. The number of type 1 and 2 outliers depends on the resolution parameter and is discussed in the choice of the resolution parameter section. The type 3 outliers were very rare after cell filtering. The supervised model evaluation below then determined the final consensus cluster version.

Supervised model evaluation on the clustering assignment. We performed a recursive feature elimination with cross-validation (RFECV)42 process from the scikit-learn package to evaluate clustering reproducibility for each consensus clustering version. We first removed the outliers from this process, and then we held out 10% of the cells as the final testing dataset. For the remaining 90% of the cells, we used tenfold cross-validation to train a multiclass prediction model using the input PCs as features and sklearn.metrics.balanced_accuracy_score43 as an evaluation score. The multiclass prediction model is based on BalancedRandomForestClassifier from the imblearn package, which accounts for imbalanced classification problems44. After training, we used the 10% testing dataset to test the model performance using the score from balanced_accuracy_score. We kept the best model and corresponding clustering assignments as the final clustering version. Finally, we used this prediction model to predict outliers’ cluster assignments. We rescued the outlier with prediction probability >0.3, otherwise labelling them as outliers.

Manifold learning for visualization. In each round of clustering analysis, the t-SNE45,46 and UMAP17 embedding were run on the PC matrix the same as the clustering input using the implementation from the scanpy40 package. The coordinates from both algorithms were in Supplementary Table 5.

Choice of resolution parameter. Choosing the resolution parameter of the Leiden algorithm is critical for determining the final number of clusters. We selected the resolution parameter by three criteria: (1). the portion of outliers <0.05 in the final consensus clustering version; (2) the ultimate prediction model accuracy >0.9; and (3) the average cell per cluster ≥ 30, which controls the cluster size to reach the minimum coverage required for further epigenome analysis such as DMR calls. All three criteria prevented the over-splitting of the clusters; thus, we selected the maximum resolution parameter under meeting the criteria using a grid search.

Cell class (level 1 clustering) annotation. We annotated non-neuron cells based on both the NeuN gate origin and low global mCH fraction. Given the strong anti-correlation between CH methylation and gene expression, we used hypo-CH-methylation at gene bodies ±2 kb of pan-excitatory markers such as Slc17a7 and Sv2b, and pan-inhibitory markers such as Gad1 and Gad2 to annotate excitatory and inhibitory cell classes, respectively.

Major type (level 2) and subtypes (level 3) annotations. We used both gene body ±2 kb hypo-CH-methylation (or hypo-CG-methylation for non-neurons) of well-known marker genes and the dissection information to annotate neuron and non-neuron clusters. All cluster marker genes are listed in Supplementary Table 7, together with the description of the cluster names, references to the marker gene information, and the URL to the data browser. The major cell types were annotated based on well-known marker genes reported in the previous studies1,19,31,47,48,49. Whenever possible, we name these clusters with canonical names (for example, IT-L23, L6b) or using descriptive names that reflect the specific spatial location of the cluster (for example, EP, CLA, IG-CA2). For subtypes, we named the clusters via its parent major type name followed by a subtype marker gene name.

Pairwise DMG identification

We used a pairwise strategy to calculate DMGs for each pair of clusters within the same round of analysis. We used the gene body ±2 kb regions of all the protein-coding and long non-coding RNA genes with evidence level 1 or 2 from the mouse GENCODE vm22. We used the single-cell level mCH fraction normalized by the global mCH level (as in ‘Computation and normalization of the methylation level’ in the clustering step above) to calculate markers between all neuronal clusters. We compared non-neuron clusters separately using the mCG fraction normalized by the global mCG level. For each pairwise comparison, we used the Wilcoxon rank-sum test to select genes with a significant decrease (hypo-methylation). Marker genes were chosen based on adjusted P < 10−3 with multitest correction using the Benjamini–Hochberg procedure, delta-normalized methylation level change <−0.5 (hypo-methylation) and area under the receiver-operating curve (AUROC) >0.8. We required each cluster to have ≥5 DMGs compared to any other cluster. Otherwise, the smallest cluster that did not meet this criterion was merged to the closest cluster based on Euclidean distance between cluster centroids in the PC matrix used for clustering. Then the marker identification process was repeated until all clusters found enough marker genes.

Three levels of iterative clustering analysis

On the basis of the consensus clustering steps described above, we used an iterative approach to cluster the data into three levels of categories. In the first level, termed CellClass, clustering analysis is done using all cells and then manually merged into three canonical classes: excitatory neurons, inhibitory neurons, and non-neurons based on marker genes. Within each CellClass, we performed all the preprocessing and clustering steps again to obtain clusters for the MajorType level using the same stop criteria. Furthermore, within each MajorType, we obtained clusters for the SubType level. All clusters’ annotations and relationships are presented in Supplementary Table 7.

Subtype taxonomy tree

To build the taxonomy tree of subtypes, we selected the top 50 genes that showed the most significant changes for each subtypes’ pairwise comparisons. We then used the union of these genes from all subtypes and obtained 2,503 unique genes. We calculated the median mCH level of these genes in each subtype and applied bootstrap resampling-based hierarchical clustering with average linkage and the correlation metric using the R package pvclust (v.2.2)50.

Impact score and total impact score

We defined the impact score (IS) to summarize pairwise comparisons for two subtype groups, where one group, A, contains M clusters and the other group, B, contains N clusters. For each gene or motif, the number of total related pairwise comparisons is M × N, the number of significant comparisons with desired change (hypo-methylation for gene or enrichment for motif) is a in group A and b in group B. The IS is then calculated as \({{\rm{IS}}}_{{\rm{A}}}\,=\,\frac{a\,-\,b}{M\,\times \,N}\) and \({{\rm{IS}}}_{{\rm{B}}}\,=\,\frac{b\,-\,a}{M\,\times \,N}\) for the two directions. For either group, IS ranges from −1 to 1, and 0 means no impact, 1 means full impact and −1 means full impact in the other group (Extended Data Fig. 7e).

We explored two scenarios using the IS to describe cluster characteristics (Extended Data Fig. 7e). The first scenario is considering each pair of branches in the subtype taxonomy tree as comprising group A and group B. Thus, the IS can quantify and rank genes or motifs to the upper nodes based on the leaves’ pairwise comparisons (Fig. 3d–f). The second scenario summarizes the total impact for specific genes or motifs regarding the taxonomy tree based on the calculation in the first scenario (Extended Data Fig. 7f–k). In a subtype taxonomy tree with n subtypes, the total non-singleton node was n − 1, and each node i had a height hi and associated ISA for one of the branches (ISB = −ISA). The node-height-weighted total IS (IStotal) was then calculated as:

$${{\rm{IS}}}_{{\rm{total}}}\,=\mathop{\sum }\limits_{i=1}^{n-1}{h}_{i}\times |{{\rm{IS}}}_{{\rm{A}}}|$$

The larger total IS indicated that a gene or motif shows more cell-type-taxonomy-related significant changes. The total IS can also be calculated in a sub-tree or any combination of interests to rank genes and motifs most related to that combination (See ‘Figure-specific methods’ for Fig. 5 regarding calculating layer and region total IS from the same tree).

Integration with snATAC-seq data

A portion of the same brain tissue sample used in this study for methylome profiling was also processed using snATAC-seq in a parallel study of chromatin accessibility3. The final high-quality snATAC-seq cells were assigned to 160 chromatin accessibility clusters (a-types). The snATAC-seq-specific data analysis steps are described in Li et al.3. Here, we performed cross-modality data integration and label-transferring to assign the 160 a-types to the 161 methylome subtypes in the following steps:

  1. (1)

    We manually grouped both modalities into five integration groups (for example, all IT neurons as a group) and only performed the integration of cells within the same group to decrease computation time. These groups were distinct in the clustering steps of both modalities and can be matched with great confidence using known marker genes. Steps 2–6 were repeated for each group. See Extended Data Fig. 5 for the group design.

  2. (2)

    We used a similar approach as described above to identify pairwise differential accessible genes (DAGs) between all pairs of a-types. The cut-off for DAG is adjusted P <10−3, fold change >2 and AUROC >0.8.

  3. (3)

    We then gathered DMGs from comparisons of related subtypes in the same group. Both DAGs and DMGs were filtered according to whether they recurred in >5 pairwise comparisons. The intersection of the remaining genes was used as the feature set of integration.

  4. (4)

    After identifying DAGs using cell-level snATAC-seq data, we merged the snATAC-seq cells into pseudo-cells to increase snATAC-seq data coverage. Within each a-type, we did a k-means clustering (k = no. of cells in that cluster/50) on the same PCs used in snATAC-seq clustering. We discarded small k-means clusters with less than 10 cells (about 5% of the cells) and merged each remaining k-means cluster into a pseudo-cell. On average, a pseudo-cell had about 50 times more fragments than a single cell.

  5. (5)

    We then used the MNN based Scanorama51 method with default parameters to integrate the snmC-seq cells and snATAC-seq pseudo-cells using genes from step 3. After Scanorama integration, we did co-clustering on the integrated PC matrix using the clustering approaches described above.

  6. (6)

    We used the intermediate clustering assignment from step 5 to calculate the overlap score (below) between the original methylome subtypes and the a-types. We used overlap score >0.3 to assign a-types to each methylome subtype. For those subtypes that have no match under this threshold, we assigned the top a-type ranked by the overlap score (Supplementary Tables 10, 11).

Overlap score

We used the overlap score to match a-type and methylome subtypes. The overlap score, range from 0 to 1, was defined as the sum of the minimum proportion of samples in each cluster overlapped within each co-cluster52. A higher score between one methylome subtype and one a-cluster indicates they consistently co-clustered within one or more co-clusters. Besides matching clusters in integration analysis, the overlap score was also used in two other cases: (1) to quantify replicates and region overlaps over methylome subtypes (Extended Data Fig. 2e–g); and (2) to quantify the overlap of each L5-ET subtype overlapping with ‘soma location’ and ‘projection target’ labels from epi-retro-seq cells (Extended Data Fig. 5j) through integration with the epi-retro-seq dataset.

Cell-type-specific regulatory elements

DMR analysis

After clustering analysis, we used the subtype cluster assignments to merge single-cell ALLC files into the pseudo-bulk level and then used methylpy (v1.4.2)38 DMRfind function to calculate mCG DMRs across all subtypes. The base calls of each pair of CpG sites were added before analysis. In brief, the methylpy function used a permutation-based root mean square test of goodness of fit to identify differentially methylated sites (DMS) simultaneously across all samples (subtypes in this case), and then merge the DMS within 250 bp into the DMR. We further excluded DMS calls that have low absolute mCG level differences by using a robust-mean-based approach. For each DMR merged from the DMS, we ordered all the samples by their mCG fraction and calculated the robust mean m using the samples between 25th and 75th percentiles. We then reassigned hypo-DMR and hyper-DMR to each sample when a region met two criteria: (1) the sample mCG fraction of this DMR is lower than (m − 0.3) for hypo-DMR or (m + 0.3) for hyper-DMR, and (2) the DMR is originally a significant hypo- or hyper-DMR in that sample judged by methylpy. DMRs without any hypo- or hyper-DMR assignment were excluded from further analyses. On the basis of these filtering criteria, we estimate the false discovery rate of calling DMRs is 2.7% (Supplementary Note 2, Extended Data Fig. 6).

Enhancer prediction using DNA methylation and chromatin accessibility

We performed enhancer prediction using the REPTILE53 algorithm. REPTILE is a random-forest-based supervised method that incorporates different epigenomic profiles with base-level DNA methylation data to learn and then distinguish the epigenomic signatures of enhancers and genomic background. We trained the model in a similar way as in the previous studies8,53, using CG methylation, chromatin accessibility of each subtype and mouse embryonic stem cells (mouse ES cells). The model was first trained on mouse ES cell data and then predicted a quantitative score that we termed enhancer score for each subtype’s DMRs. The positives were 2 kb regions centred at the summits of the top 5,000 EP300 peaks in mouse ES cells. Negatives include randomly chosen 5,000 promoters and 30,000 2-kb genomic bins. The bins have no overlap with any positive region or gene promoter8.

Methylation and chromatin accessibility profiles in bigwig format for mouse ES cells were from the GEO database (GSM723018). The mCG fraction bigwig file was generated from subtype-merged ALLC files using the ALLCools package ( For chromatin accessibility of each subtype, we merged all fragments from snATAC-seq cells that were assigned to this subtype in the integration analysis and used deeptools bamcoverage to generate CPM normalized bigwig files. All bigwig file bin sizes were 50 bp.

Motif-enrichment analysis

We used 719 motif PWMs from the JASPAR 2020 CORE vertebrates database54, where each motif was able to assign corresponding mouse transcription factor genes. The specific DMR sets used in each motif-enrichment analysis are described in figure specific methods below. For each set of DMRs, we standardized the region length to the centre ±250bp and used the FIMO tool from the MEME suite55 to scan the motifs in each enhancer with the log-odds score P <10−6 as the threshold. To calculate motif enrichment, we use the adult non-neuronal mouse tissue DMRs10 as background regions unless expressly noted. We subtracted enhancers in the region set from the background and then scanned the motifs in background regions using the same approach. We then used Fisher’s exact test to find motifs enriched in the region set and the Benjamini–Hochberg procedure to correct multiple tests. We used the TFClass56 classification to group transcription factors with similar motifs.

DMR–DMG partial correlation

To calculate DMR–DMG partial correlation, we used the mCG fraction of DMRs and the mCH fraction of DMGs in each neuronal subtype. We first used linear regression to regress out variance due to global methylation difference (using scanpy.pp.regress_out function), then use the residual matrix to calculate the Pearson correlation between DMR and DMG pairs where the DMR centre is within 1 Mb of the TSSs of the DMG. We shuffled the subtype orders in both matrices and recalculated all pairs 100 times to generate the null distribution.

Identification of loops and differential loops from sn-m3C-seq data

After merging the chromatin contacts from cells belonging to the same type, we generated a .hic file of the cell-type with Juicer tools pre. HICCUPS57 was used to identify loops in each cell type. The loops from eight major cell types were concatenated and deduplicated and used as the total samples for differential loop calling. A loop-by-cell matrix was generated, in which each element represents the number of contacts supporting each loop in each cell. The matrix was used as input of EdgeR to identify differential interactions with ANOVA tests. Loops with FDR <10−5 and minimum–maximum fold change >2 were used as differential loops. Note that the abundance of cell types is highly variable, leading to different coverages of contact maps after merging all the cells from each cell type. Since HICCUPS loop calling is sensitive to the coverage, more loops were identified in the abundant cell types (for example, 12,614 loops were called in DG, containing 1,933 cells) compared to the less abundant ones (for example, 1,173 loops were called in MGE, containing 145 cells). Therefore, we do not compare the feature counts related to the loops across cell types directly in our analyses.

Figure-specific methods

3D model of dissection regions (Fig. 1b–e). We created in silico dissection regions based on the Allen CCF16 3D model using Blender 2.8 that precisely follow our dissection plan. To ease visualization of all different regions, we modified the layout and removed some of the symmetric structures, but all the actual dissections were applied symmetrically to both hemispheres.

Calculating the genome feature detected ratio (Extended Data Fig. 2a). The detected ratio of chromosome 100-kb bins and gene bodies is calculated as the percentage of bins with >20 total cytosine coverage. Non-overlapping chromosome 100-kb bins were generated by bedtools makewindows -w 100000; gene bodies were defined by GENCODE vm22.

Integration with epi-retro-seq L5-ET cells (Fig. 2g–j, Extended Data Fig. 5g–j). Epi-retro-seq is an snmC-seq2-based method that combines retrograde AAV labelling22. The L5-ET cells’ non-overlapping chromosome 100-kb bin matrix gathered by the epi-retro-seq dataset was concatenated with all the L5-ET cells from this study for co-clustering and embedding as described in ‘Clustering-related methods’. We then calculated the OS between subtypes in this study and the ‘soma location’ or ‘projection target’ labels of epi-retro-seq cells. The first OS helped quantify how consistent the spatial location is between the two studies; the second OS allowed us to impute the projection targets of subtypes in this study.

Pairwise DMR and motif-enrichment analysis (Fig. 3c, f). The total subtype DMRs were identified as described in ‘Cell-type-specific regulatory elements’ by comparing all subtypes. We then assigned DMRs to each subtype pair if the DMRs were: (1) significantly hypomethylated in only one of the subtypes; and (2) the mCG fraction difference between the two subtypes is >0.4. Each subtype pair was associated with two exclusive sets of pairwise DMRs. We carried out motif-enrichment analysis described in ‘Cell-type-specific regulatory elements’ on each DMR set using the other set as background. Motifs enriched in either direction were then used to calculate the impact score and were associated with upper nodes of the taxonomy.

Overlapping eDMR with genome regions (Fig. 4b). The cluster-specific snATAC-seq peaks were identified in Li et al.3. We used bedtools merge to aggregate the total non-overlap peak regions and bedtools intersect to calculate the overlap between peaks and eDMRs. The developing forebrain and other tissue feDMRs were identified in He et al.8 using methylC-seq58 for bulk whole-genome bisulfite sequencing. All of the genome features used in Fig. 4b were defined as in He et al.3, except using an updated mm10 CGI region and RepeatMaster transposable elements lists (UCSC table browser downloaded on 9 October 2019).

Heat maps of the gene–enhancer landscape (Extended Data Fig. 8e). The eDMRs for each gene were selected by eDMR–gene correlation of >0.3. Sections of the heat maps in Extended Data Fig. 8e were gathered by (1) mCG fraction of each eDMR in 161 subtypes from this study; (2) snATAC-seq subtype-level fragments per kilobase of transcript per million mapped reads (FPKM) of each eDMR in the same subtype orders. The subtype snATAC profiles were merged from integration results as described in ‘Clustering-related methods’; (3) mCG fraction of each eDMR in forebrain tissue during ten developing time points from embryonic day 10.5 (E10.5) to P0 (data from He et al.8); (4) H3K27ac FPKM of each eDMR in 7 developing time points from E11.5 to P0 (data from Gorkin et al.59); (5) H3K27ac FPKM of each eDMR in P56 frontal brain tissue (data from Lister et al.6); and (6) eDMR is overlapped with forebrain feDMR using bedtools intersect.

Embedding of cells with chromosome interactions (Fig. 4e). scHiCluster60 was used to generate the t-SNE embedding of the sn-m3C-seq cells. Specifically, a contact matrix at 1-Mb resolution was generated for each chromosome of each cell. The matrices were then smoothed by linear convolution with pad = 1 and random walk with restart probability = 0.5. The top 20th percentile of strongest interactions on the smoothed map was extracted, binarized and used for PCA. The first 20 PCs were used for t-SNE.

IT layer dissection region group DMG and DMR analysis (Fig. 5a–c). To collect enough cells for dissection region analysis, we used only the major types (corresponding to L2/3, L4, L5 and L6) of IT neurons. We grouped cells into groups according to layer dissection region and kept groups with >50 cells for further analysis (Extended Data Fig. 9b). We performed pairwise DMG, DMR and motif-enrichment analysis, the same as the subtype analysis in Fig. 3, but using the layer dissection region group labels. We then built a spatial taxonomy for these groups and used it to calculate impact scores. To rank layer-related or dissection-region-related genes and motifs separately, we used two sets of the branches (Extended Data Fig. 9c, top set for layers, bottom set for regions) in the taxonomy and calculated two total impact scores using the equations above.

DG cell group and gradient DMR analysis (Fig. 5e). DG cells were grouped into four evenly sized groups according to the cells’ global mCH levels, with cut-off thresholds at 0.45%, 0.55% and 0.69%. We then randomly chose 400 cells from each group to call gradient-DMRs using methods described in ‘Clustering-related methods’. To ensure the DMRs identified between intra-DG groups were not due to stochasticity, we also randomly sampled 15 groups of 400 cells from all DG cells regardless of their global mCH and called DMRs among them as control-DMRs (2,003 using the same filtering condition). Only 0.04% of gradient DMRs overlapped with the control DMRs; these were removed from further analysis. Pearson correlations (ρ) of mCG fractions of each gradient DMR was calculated against a linear sequence (1, 2, 3, 4) to quantify the gradient trend. DMRs with ρ <−0.75 or ρ >0.75 were considered to be significantly correlated. Weakly correlated DMRs (10% of DMRs) were not included in further analysis.

DMR- and DMS-enriched genes (Fig. 5f, g). To investigate the correlated DMR or DMS enrichment in specific gene bodies, we compared the number of DMS and cytosine inside the gene body with the number of DMS and cytosine in the ±1 Mb regions using Fisher’s exact test. We chose genes passing two criteria: (1) adjusted P <0.01 with multitest correction using the Benjamini–Hochberg procedure, and (2) overlap with >20 DMSs. Gene ontology analysis of DMR and DMS enriched genes was carried out using GOATOOLS61. All protein-coding genes with gene body length >5 kb were used as background to prevent gene-length bias.

Compartment strength analysis (Extended Data Fig. 10g). We normalized the total chromosome contacts by z-score in each 1-Mb bin of the DG contact matrix, and the bins with normalized coverage between −1 and 2 were kept for the analysis. After filtering, the PC1 of the genome-wide Knight–Ruiz-normalized62 contact matrix was used as the compartment score. The score was divided into 50 categories with equal sizes from low to high, and bins were assigned to the categories. The intra-chromosomal observation/expectation (ove) matrices of each group were used to quantify the compartment strength. We computed the average ove values within each pair of categories to generate the 50 × 50 saddle matrices. The compartment strength was computed with the average of the upper left and lower right 10 × 10 matrices divided by the average of the upper right and lower left 10 × 10 matrices63.

Domain analysis (Extended Data Fig. 10i). We identified 4,580 contact domains at 10-kb resolution in DG using Arrowhead57. For bin \(i\), the insulation score I is computed by

$${I}_{i}=\frac{{{\rm{m}}{\rm{e}}{\rm{a}}{\rm{n}}}_{i-10\le {i}^{{\prime} } < i;i\le {j}^{{\prime} } < i+10}{A}_{{i}^{{\prime} }{j}^{{\prime} }}}{max({{\rm{m}}{\rm{e}}{\rm{a}}{\rm{n}}}_{i-10\le {i}^{{\prime} } < i;i-10\le {j}^{{\prime} } < i}{A}_{{i}^{{\prime} }{j}^{{\prime} }},{{\rm{m}}{\rm{e}}{\rm{a}}{\rm{n}}}_{i\le {i}^{{\prime} } < i+10;i\le {j}^{{\prime} } < i+10}{A}_{{i}^{{\prime} }{j}^{{\prime} }})}$$

where A is the ove of Knight–Ruiz-normalized matrices and mean is the average of A over the range in the subcript. For each group, insulation scores of domain boundaries and 100-kb flanking regions were computed and averaged across all boundaries.

Prediction model description

Related to Fig. 6. To reduce the computing complexity, we applied PCA on the dataset of 100-kb bin mCH features to obtain the first 3,000 PCs, which retain 61% of the variance of the original data. These 3,000 PCs were then used to train and test the predicting model. We used an ANN with two hidden layers to simultaneously predict cell subtypes and their dissection regions. The input layer contains 3,000 nodes, followed by a shared layer with 1,000 nodes. The shared layer is further connected simultaneously to two branch hidden layers of the subsection region’s subtype, each containing 200 nodes. The corresponding one-hot encoding output layers follow branch hidden layers. We used fivefold cross-validation to access the model performance. We applied the dropout technique64 with a dropout rate P = 0.5 on each hidden layer to prevent overfitting during the training. Adam optimization65 was used to train the network with a cross-entropy loss function. The training epoch number and batch size are 10 and 100, respectively. The training and testing processes were conducted via TensorFlow 2.066.

Model performance. The two output layers generate two probabilistic vectors for each single cell input as the prediction results for cell subtypes and dissection regions, respectively. The subtype and dissection region label with the highest probabilities were used as the prediction results for each cell to calculate accuracy. When calculating the cell dissection region accuracy (Fig. 6c), we defined two kinds of accuracy with different stringency: (1) the exact accuracy using the predicted label, and (2) the fuzzy accuracy using predicted labels or its potential overlap neighbours. The potential overlap neighbours curated based on Allen CCF (Extended Data Fig. 11a, Supplementary Table 2) stood for adjacent regions of a particular dissection region. The exact accuracy of the ANN model is 69% and the fuzzy accuracy is 89%. To evaluate how much of the dissection region accuracy was improved via ANN, we calculated fuzzy accuracy based only on naive guesses in each subtype based on the dissection region composition (grey dots in Extended Data Fig. 11c). We also trained additional models using logistic regression and random forest for benchmarks. The performance of ANN on subtype prediction is comparable with logistic regression and random forest. By contrast, the performance in location prediction is substantially improved against the other two models (Extended Data Fig. 11b), suggesting that distinguishing the cells from different dissected regions may require nonlinear relationships between genomic regions. We used scikit-learn (v0.23) for logistic regression and random forest implementation and the multinomial objective function for multi-class classification. N_estimators were set to 1,000 for the random forest.

Biological feature importance for dissection region prediction (Fig. 6e). To assess which DNA regions store information of cell spatial origins that is distinguishable using our model, we evaluated the importance of PC features by examining how permutation of each PC feature across cells affects prediction accuracy. We tested five permutations for each feature and used decreasing average accuracy to indicate PC feature importance. We examined genes contained in the 100-kb bins with the top 1% PCA factor loadings for the most important PC feature for a given cell type.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this paper.