Tissue-specific DamID protocol using nanopore sequencing

Georgina Gómez-Saldivar1, Dominique A. Glauser1*, Peter Meister2*
1Department of Biology, University of Fribourg, 1700 Fribourg, Switzerland
2Cell Fate and Nuclear Organization, Institute of Cell Biology, University of Bern, 3012 Bern, Switzerland
*Corresponding authors: Dominique A. Glauser, Email:; Peter Meister, Email:
Competing interests: The authors have declared that no competing interests exist.
Abbreviations used: ChIP, chromatin immuno-precipitation; Dam, DNA adenine methyltransferase; DamID, DNA adenine methylation identification; ONT, Oxford Nanopore technologies; POI, protein of interest; RAPID, RNA PoI DamID; FACS, fluorescence activated cell sorting; FANS, fluorescence activated nuclei sorting; SRT, trans-splicing-based RNA tagging; PAT-seq, polyA tagging and sequencing; gDNA, genomic DNA; ds-adapters, double-stranded adapters; SQB, sequencing buffer; LB, loading beads; FLT, flush tether; FB, flush buffer; GFF, general feature format; RPB-6, RNA polymerase subunit 6; LMN-1, lamin; GFP, green fluorescent protein
Received March 23, 2021; Revision received May 5, 2021; Accepted May 12, 2021; Published August 27, 2021
DNA adenine methylation identification (DamID) is a powerful method to determine DNA binding profiles of proteins at a genomic scale. The method leverages the fusion between a protein of interest and the Dam methyltransferase of E. coli, which methylates proximal DNA in vivo. Here, we present an optimized procedure, which was developed for tissue-specific analyses in Caenorhabditis elegans and successfully used to footprint genes actively transcribed by RNA polymerases and to map transcription factor binding in gene regulatory regions. The present protocol details C. elegans-specific steps involved in the preparation of transgenic lines and genomic DNA samples, as well as broadly applicable steps for the DamID procedure, including the isolation of methylated DNA fragments, the preparation of multiplexed libraries, Nanopore sequencing, and data analysis. Two distinctive features of the approach are (i) the use of an efficient recombination-based strategy to selectively analyze rare cell types and (ii) the use of Nanopore sequencing, which streamlines the process. The method allows researchers to go from genomic DNA samples to sequencing results in less than a week, while being sensitive enough to report reliable DNA footprints in cell types as rare as 2 cells per animal.
Graphic Abstract
Keywords: CRE recombinase-based, tissue-specific expression, DNA adenine methylation identification, DNA-protein interaction, long-read sequencing, third generation sequencing


The advent of deep sequencing has revolutionized basic and applied biomedical science. Analyzing the expression of an entire genome, how its expression is modified by intra- and extra-cellular factors, or obtaining the genomic footprints of DNA-binding proteins have become routine procedures in many research projects. However, when working with entire animals, evaluating genome-wide changes for a given cell type often remains challenging, as it requires to either sort cells from a dissociated tissue of interest or use transgenic approaches to purify transcripts from the target cell type [1-4]. This can be difficult when the considered cell type is hard to separate from surrounding cells or is a rare cell type. In particular, in the nematode C. elegans for which no continuous culture system for differentiated cells has been developed, this is a major hurdle to identify expressed genes or determine interaction profiles of DNA-binding proteins in rare cell types.

DNA adenine methylation identification

DNA adenine methylation identification (DamID) [5] was developed in 2000 by Bas van Steensel and Steven Henikoff as an alternative to chromatin immuno-precipitation (ChIP) to identify genome-wide DNA-protein interactions in vivo, avoiding the troublesome steps of cell or nucleus isolation, fixation and immunoprecipitation with antibodies. DamID relies on the expression of trace levels of the E. coli DNA adenine methyltransferase (Dam) fused to a protein of interest (POI). The Dam methylates DNA located close to the binding site of the POI at GATC sites (Fig. 1A). Methylated sites are then used to track POI binding sites. Notably, DamID is highly versatile and has been applied in many organisms, ranging from fission yeast [6-10], C. elegans [11-17], Drosophila [18-23], Medaka [24], Arabidopsis [25-27], and mice [28-31] to human cells [32-35]. Additionally, DamID has been adapted to evaluate an extensive range of chromatin features in vivo such as chromatin accessibility (CATaDA) [36], chromatin dynamics visualization (m6A-tracer) [37], transcription factor binding (MaTaDa) [38] and co-binding (SpDamID) [39], RNA-DNA interactions (RNA-DamID) [40], three-dimensional genome organization (DamC) [41], transcriptional profiling (TaDa [42] and RAPID [43]), or simultaneous transcription and protein-DNA interaction profiling in single cells (scDam&T-seq) [44,45].

Development of tissue-specific DamID in C. elegans

We recently developed RAPID (RNA PoI DamID) to profile genome-wide, tissue-specific RNA Polymerase occupancy in vivo without cell isolation or nuclei purification [43]. In contrast to other methods, which sequence transcripts from specific cell types, RAPID is based on the methylation footprinting by an RNA polymerase subunit fused to a Dam methyltransferase, expressed only in the target tissue (Fig. 1B and 1C). Using DNA purified from entire animals, we profiled muscle and intestine transcription, with results comparable to other tissue-specific RNA-seq methods based on sorting (Fluorescence Activated Cell or Nuclei Sorting: FACS [1], FANS [2]) or biochemical purification (trans-splicing-based RNA tagging, SRT [3], and polyA tagging and sequencing, PAT-seq [4]). We further showed that this method can be applied for rare cell types by profiling cell types representing only two cells per animal or 0.2% of the somatic cells. The specific advantages and limitations of RAPID as compared to RNA-seq-based methods were extensively discussed in this previous work [43].
Figure 1. Tissue-specific DamID schemes. A. DamID is used to identify genome-wide DNA-protein interactions in vivo. DNA adenine methyltransferase (Dam) fused to a protein of interest (POI, here a subunit of the RNA polymerase) methylates GATC sites, forming unique flags to track genome regions where the POI was associated. B. Recombination-based transgenic approach for tissue-specific DamID. Tissue-specific DamID is achieved by in vivo CRE recombination in the tissue of interest, deleting a floxed mCherry::STOP cassette and enabling the transcription of the Dam::POI fusion or the Dam::GFP fusion, respectively. The later one is used to normalize for overall chromatin accessibility. Red arrows represent primers used to verify CRE recombination (see Fig. 4). C. Illustration of tissues already successfully used with tissue-specific DamID representing from 10% to 0.2% of the somatic cell content of the animal.
Although we applied this approach to RNA polymerase, the same approach can additionally be applied to profile the genome-wide interactions of any POI in a tissue-specific manner. Tissue-specific profiling requires only a modest number of animals (4000–5000), thereby reducing the time required to grow biological sample populations. Additionally, we coupled the methylation footprinting method with long-read sequencing technology using Oxford Nanopore technologies (ONT) [46]. This further reduces the data acquisition time, because sequencing can be performed in every laboratory without the requirement for a sequencing facility. Here, we present a bench protocol allowing one to perform methyl footprinting from DNA extraction to amplicon sequencing in less than a week.

Tissue-specific DamID in C. elegans

DamID requires the expression of a Dam fusion protein, which, if expressed at moderate to high levels, could in some instances lead to toxicity and potentially to mutations (data not shown) [42]. This is solved in C. elegans by the use of inducible promoters in the uninduced state to achieve very low expression levels. These promoters are however broadly expressed and therefore unadapted for tissue-specific profiling. In our tissue-specific DamID approach, the Dam-POI expression is driven by a heat shock promoter flanked in 5’ by a loxP-mCherry-STOP-loxP cassette impairing its expression in non-target tissues (Fig. 1B). An additional construct drives the CRE recombinase expression in a cell-type specific manner, excising the STOP-cassette and thereby allowing basal expression of the Dam-POI. DamID requires a control for chromatin accessibility, as methylation by the POI Dam fusion depends on it. As a control for overall chromatin accessibility, we use a similar mCherry floxed GFP-Dam construct, which is expressed in the same tissue in a control line treated in parallel.

Critical aspects to consider before performing tissue-specific DamID

DamID has many advantages, including its very low background noise and exquisite sensitivity. However, before embarking on the use of tissue-specific DamID, it is essential to be aware of its limits.
Steric effects of Dam fusion: One of the initial points to consider is whether the Dam should be fused at the N- or C-terminus of the POI, in order to minimize the steric effect of the Dam and its impact on POI functionality, including its DNA-binding ability. This choice could be potentially guided by pre-existing information on the functional or binding domains of the POI, or by data from previous experiments in which fusion protein constructs have been functionally evaluated. Even if a systematic comparison of N- and C-terminal fusion designs on different transcription factors has shown robust target footprints regardless of the Dam position, the potential Dam steric effect remains hard to predict and may vary on a case-by-case basis [47].
Spatial resolution: DamID spatial resolution depends on the density of GATC sites in the genome. C. elegans has 269049 GATC motifs per haploid genome [48], corresponding to a mean distance between sites of 374 bp and a median of 210 bp [49]. With this density, DamID was successfully applied to obtain the genomic footprint of several transcription factors such as DAF-16 [13], CEH-60 [14] and CRH-1 (our unpublished data), as well as RNA Polymerases [43]. However, for most genomic regions the binding sites are defined as relatively large windows of several hundred bases, sometimes much larger in some regions with low GATC density. This corresponds to a spatial resolution substantially lower than a typical interval determined by ChIP.
Methylation dynamics: Understanding the methylation dynamics is essential to plan and interpret DamID experiments. Three aspects are particularly important. First, the irreversibility of DNA methylation in C. elegans, which lacks any relevant DNA demethylase activity. In a non-replicating C. elegans cell, methylation marks will therefore be irreversibly deposited. Second, the impact of DNA replication. Methyl marks will stay until DNA replication occurs and one DNA strand is passively demethylated through the DNA replication process. Hemimethylated DNA is not cut by DpnI and will remain undetected. Third, considering cell populations, the average methylation at a given site will build up over time. If Dam-POI interactions are transient or intermittent the level of methylation is expected to increase more slowly than for robust, long-lasting interactions. Consistent with these theoretical considerations, the profiles obtained in post-mitotic cells of adult worms tend to be cleaner and with better correlation between replicates compared to those obtained in embryos [11,43], presumably because the Dam-POI fusion protein has been expressed for a very short time in the early stages of development with fast-replicating cells. Eventually, a good knowledge about the timing of the cell lineage replication and the CRE transgene expression onset is essential to plan DamID experiments and interpret their results.


Construction of DamID tissue-specific strains

Nematode culture

Purification of gDNA

Digestion and purification of methylated DNA

Multiplexed library preparation for DamID nanopore sequencing

NOTE: This module is part of the Ultra™ II workflow and is compatible for library construction in both Illumina® and Oxford Nanopore Technologies® workflows.
NOTE: Native Barcoding Expansion, is a kit to multiplex samples using PCR-free method to preserve additional information such as base modifications. Using both expansion kits (EXP-NBD104 and EXP-NBD114), there are 24 unique barcodes, allowing the user to pool up to 24 different samples in one sequencing experiment.


The protocol presented below is visually summarized in Figure 2.
Figure 2. Overview of the entire tissue-specific DamID protocol. The scheme is divided into four main parts: I. Construction of DamID tissue-specific plasmids and strains (step 1). II. Worm harvesting (step 2) and extraction of gDNA as starting material (step 3). III. Amplification of methylated fragments (step 4), purification with magnetic beads (represented with a red magnet), and quantification with Qubit (represented with a blue square) and preparation of multiplex libraries (step 5). IV. Sequencing of libraries using Nanopore technology (step 6) and data analysis (step 7).

1.Construction of DamID tissue-specific strains

The tissue-specific promoter driving the expression of CRE protein (promoter::CRE) is selected according to the tissue to be studied (like for the examples in Fig. 1C). The CRE expression plasmid can be easily designed with Gibson Assembly technology, allowing the fast and easy analysis of a POI in multiple tissues (Fig. 3).
NOTE: Pre-existing CRE expression lines containing single copy transgenes (Fig. 3 [43,51]) or integrated transgene arrays [52] could potentially be used.
The plasmid expressing Dam-POI should be designed for each protein to analyse, whereas the GFP-Dam vector, pCFJ150 hsp-16.2p::loxP::mCherry::STOP::loxP::gfp::dam::unc-58 3’ UTR [43], remains constant. Both plasmids contain the loxP::mCherry::STOP::loxP cassette.

1.1.Using standard molecular cloning techniques clone the gene of interest into a Dam expression plasmid of choice, e.g., plasmid #503 [pCFJ150 hsp-16.2p::loxP::mCherry::STOP::loxP::dam::rpb-6::unc-54 3’ UTR] or #418 [pCFJ150 hsp-16.2p::loxP::mCherry::STOP::loxP::gfp::dam::unc-54 3’ UTR], depending on whether N- or C-terminal tagging is preferred [43,47]. Annotated plasmid sequences are provided as supplementary material (File S1 and S2). The transgene must be under the control of an inducible promoter with low basal activity and a characterized 3′UTR. The plasmids above include the hsp-16.2 promoter and the unc-54 3’UTR (Fig. 3A).

1.2.Verify the sequence using primers that cover the insertion sites and the gene of interest. With the plasmids above, primers Dam-forward and unc-54-reverse can be used.

1.3.Using standard molecular cloning techniques, such as Gibson Assembly, clone the promoter of interest into a CRE expression plasmid of choice, e.g., pSR33 (Fig. 3B [51]).

Figure 3. Design of the plasmids used in tissue-specific DamID. A. Outline of the Dam::POI (protein of interest) plasmid construction using Gibson Assembly technology. Blue arrows represent the Gibson Assembly primers that amplify the gene of interest, violet arrows represent the Gibson Assembly primers that amplify the dam gene. Scissors represent restriction sites for excision of gfp:dam fragment in pCFJ150 plasmid. Using Gibson Assembly reaction, the gene of interest (blue) and the dam (violet) fragments are inserted in the pCFJ150 backbone. B. Outline of the tissue-specific CRE expression plasmid construction using Gibson Assembly technology. Red arrows represent the Gibson Assembly primers to amplify the tissue-specific promoter of interest. Scissors represent restriction sites for excision of hsp-16.2p in pSR33 plasmid. Using a Gibson Assembly reaction, a new promoter fragment (blue) is inserted in the pSR33 backbone.

1.4.Verify the sequence of the insertion. In the pSR33 backbone, M13 forward and M13 reverse primers can be used for sequencing. Depending on the size of the inserted promoter, additional sequencing primers may need to be designed.

1.5.Inject the DamID vectors into the gonad of an appropriate C. elegans host strain for MosSCI integration and isolate transgenic lines according to standard MosSCI protocols [50]. Dam-POI constructs are integrated either on chromosome II or IV and promoter::CRE constructs are integrated on chromosome X.

1.6.When designing new CRE drivers to target specific cell types, verify the correct localization and recombination function of CRE in vivo, by crossing the promoter::CRE line with the reporter line SV1361 available from CGC [51]. This reporter line ubiquitously expresses a nuclear red marker from the [rps-27p::loxP::NLS::mCherry::let858 3'UTR::loxP::NLS::GFP::let-858 3'UTR] transgene. Upon successful recombination of the mCherry cassette, CRE-expressing cells will start to produce NLS::GFP, labeling the corresponding nuclei in green (Fig. 4B).

1.7.Cross the strains expressing Dam-POI and promoter::CRE to generate a line being homozygous for both transgenes.

1.8.Using PCRs, verify the Dam-POI recombination causing the loxP::mCherry::STOP::loxP cassette excision (Fig. 4A). The primers design must ensure that two amplification products are distinguished (Fig. 1B): a small one, corresponding to the genomic DNA from the CRE-expressing tissues, for which the mCherry cassette is excised; and a large one, corresponding to the rest of the tissues, in which CRE is absent and the mCherry cassette is still present (Fig. 4A).

Figure 4. Verification of CRE-mediated recombination. A. Verification of recombination using PCR. Representative agarose gel electrophoresis results, showing three successfully recombined samples (left lanes), a size ladder, and four controls. Primers should be designed to evaluate the presence or the absence of loxP::mCherry::STOP::loxP cassette (as schematized in Fig. 1B). A successful PCR should generate two amplification products: a small one (about 500 bp), corresponding to the gDNA from the CRE-expressing tissues for which the mCherry cassette is excised; and a large one (about 1700 bp), corresponding to the rest of the tissues in which the mCherry cassette is still present. Note: Although a smaller number of cells are expressing the recombined product, the smaller PCR product tends to be more abundant due to PCR bias favoring the amplification of smaller products. B. Verification of CRE-mediated recombination in the selected tissue with a reporter transgene. The depicted example shows the CRE expression in XXX neuroendocrine cells in vivo. An XXX-specific CRE driver strain using the sdf-9p promoter [DAG827; 43] was crossed with the reporter strain SV1361, which carries a floxed mCherry::STOP cassette in front of an nls::gfp open reading frame. Upon successful recombination, the mCherry cassette is excised and CRE-expressing cells produce NLS::GFP, here labeling XXX neuroendocrine nuclei (white arrow). The asterisks mark autofluorescence of intestinal cells.

2.Nematode culture

2.1.Collect embryos from asynchronous cultures by standard hypochlorite treatment.

2.1.1.Wash 3–4 mixed-stage 6 cm NGM plates containing many gravid adults with M9 and transfer to a 15 ml Falcon tube.

2.1.2.Centrifuge at 400 g for 2 min in a swinging bucket centrifuge, and discard the supernatant using a vacuum pump.

2.1.3.Wash animals twice with 14 ml M9 buffer. Each time, centrifuge at 400 g for 2 min and discard the supernatant. Repeat if the solution is still cloudy, a sign that bacteria are still present.

2.1.4.Add 2 ml of fresh bleach solution at room temperature (recipe for 5 ml: 3.5 ml Milli-Q water, 1 ml 13% sodium hypochlorite, 0.5 ml 5M KOH). Agitate the tubes by hand and vortex occasionally (no more than 4 min; and no more than 4–6 strains at a time). Check worms regularly with a dissecting stereoscope. When 60% of the worms are broken up, proceed to the next step. In subsequent steps, work under sterile conditions to avoid contaminations with Dam positive bacteria.

2.1.5.Add 13 ml Milli-Q water and completely resuspend the pellet to avoid worm death.

2.1.6.Centrifuge at 800 g for 1 min and discard supernatant.

2.1.7.Wash 5 times with 13 ml M9, centrifuge at 800 g for 1 min and discard supernatant. After the last wash, resuspend in 5 ml of M9. Count the number of embryos in 2–10 μl aliquots.

2.1.8.Leave embryos to hatch overnight (14–16 h) at 16–20°C with gentle agitation on a roller.

2.1.9.Centrifuge at 600 g for 1 min and discard supernatant leaving about 1–1.5 ml to resuspend the L1 larvae.

2.2.Count hatched L1s in 2–10 μl aliquots.

2.3.Place 1500–2000 L1 animals per 10 cm plate containing a thick lawn of Dam negative E. coli bacteria (e.g., strain GM48, GM119) as a food source (prepared with 1 ml of a fresh Dam- overnight liquid culture). Set at least two cultures per strain, as biological replicates.

2.4.Incubate worms at 15°C to 20°C and let them grow for two generations, making sure they never run out of food.

2.5.Once the plate contains mainly gravid adults, wash the plates with M9 and transfer the liquid to a 15 ml falcon tube.

2.6.Repeat steps 2.1 and 2.2.

NOTE: The first bleaching is to remove traces of OP50 Dam positive E. coli. This strain would create a high background Dam signal as methylated bacterial DNA is co-purified with the worm DNA. Animals are then grown with GM48 Dam negative E. coli. We estimate that after 2 generations most remaining bacterial DNA is not methylated. The second bleaching is to synchronize worm growth for the experiment.

2.7.Place 1000–1250 L1s per 10 cm plate containing a thick lawn of Dam negative E. coli bacteria (prepared with 1 ml of fresh Dam negative strain grown overnight in LB and left to dry for 5 h to overnight once spread on the plate). Use a total of four 10 cm-plates per strain and per biological replicate, in order to grow a total of 4000–5000 worms for each replicate.

2.8.Incubate worms at 15°C to 20°C and collect animals when they reach the desired life stage (e.g., 74 h for L4; 94 h for non-gravid young adults, etc.).

2.9.Harvest worms and wash them 5 times with 15 ml M9 to remove bacteria. Do not centrifuge, but let worms go down by decantation; later wash worms 5–7 more times with M9 and centrifugation at 400 g for 1 min. Keep the M9 at 15°C to avoid heat-shocking the animals.

2.10.After the last wash, resuspend the worms in less than 1.5 ml M9 and transfer the suspension to 1.5 ml microcentrifuge tubes. Centrifuge at 13000 g for 30 s, discard supernatant and make aliquots containing about 30–35 μl worm material. Remove as much as possible the excess liquid to obtain a worm pellet of about 30 mg, snap-freeze in liquid nitrogen and store at −80°C until needed.

Figure 5. Standardization of DamID PCR. Examples of agarose gel electrophoresis analyses of amplified methylated fragments in tissue-specific DamID samples and required controls (A, sample with methylated DNA, without DpnI but with ligase; B, sample with methylated DNA, with DpnI but without ligase; C, sample with water, DpnI and ligase). The appropriate number of PCR cycles used to amplify the methylated fragments for each Dam-POI has to be determined according to the tissue where it is expressed. A. Agarose gel showing the smear signal obtained for Dam::RBP-6 in XXX cells over a range of 20 to 30 cycles. Several controls (A, B and C) are analyzed in parallel to the samples to rule out the production of non-DamID amplicons. The cycle number selection is based on a strong signal for the sample (complete reaction: methylated DNA, DpnI and ligase) and an absence of signal in the controls. B. Same as in A for muscle cells. One asterisk indicates the selected number of cycles for each tissue (24 for XXX cells; 22 cycles for muscle cells). The cycle number selection is based on a strong signal for the sample and an absence of signal in the controls (A, B, C, and N2: sample of unmethylated DNA, DpnI and ligase). Two asterisks indicate the cycle where a non-specific signal starts to be visible for one or more of the controls. These might arise from contamination with methylated E. coli DNA (control A), gDNA breaks (control B) or contaminated reagents (control C or water), as well as from non-specific amplicons arising from non-methylated worm DNA (control N2). C. Agarose gel showing amplicons obtained from DNA of strains expressing different Dam-POI in two different cell types (each time two biological replicates). Left: amplicons obtained from strains with intestine-specific expression of Dam fusions to RPB-6 (RNA polymerase subunit 6), GFP (chromatin accessibility control) and LMN-1 (lamin). The RNA polymerase subunit(RBP-6) produces stronger signals than LMN-1, associated with a shorter proportion of the genome. Right: amplicons obtained from strains with XXX-specific expression of Dam fusions to RPB-6 (RNA polymerase), GFP (chromatin accessibility control) and CRH-1 (a transcription factor). The RNA polymerase subunit (RPB-6) produces a stronger DamID signal than the CRH-1 transcription factor. D. Example of successful amplification of Dam-methylated DNA after standardizations. Dam-POI and GFP-Dam experiments were processed and run in parallel with all technical controls. Only a small volume of the preparation was analyzed on gel for this verification and the rest could be used for the DamID experiment. Methylated DNA amplicons are visible as a smear (400–1200 bp range).

4.Digestion and purification of methylated DNA

4.1.Digest 500 ng gDNA using 10 units DpnI (NEB #R0176S; cuts methylated GmATC).

4.1.1.Set up DpnI digestion reactions on ice (Table 1). Make sure to prepare additional tubes for controls: (i) without DpnI (one control for each biological sample); (ii) for one of the biological samples with methylated DNA, a second digestion replicate control digested with DpnI, which will be later used without ligase (Fig. 5A).

Table 1. DpnI digestion mix.
Component Volume per reaction (μl) Final
10× CutSmart 1
DpnI (20 U/μl) 0.5
gDNA X (max. 8.5 µl) 500 ng

4.1.2.Incubate 6 h at 37°C in a thermocycler.

4.1.3.Heat-inactivate the enzyme at 80°C for 20 min and cool at 4°C.

4.2.Prepare double-stranded adapters (ds-adapters, each at 50 μM final concentration).

4.2.1.Mix 50 μl of primer AdRt (100 μM) and 50 μl of AdRb (100 μM).

4.2.2.Anneal the primers in a thermocycler, using the following program: 95°C for 10 min and then reduce 5 degrees every 1 min until 25°C, then 12°C forever (at least 10 min).

Note: The ds-adapter mix can be prepared in advance and kept frozen at −20°C.

4.3.Ligate the adapters to DpnI-digested DNA

4.3.1.Prepare the ds-adapter ligation reaction mix on ice (Table 2), as well as the control B (see above) with DpnI-digested DNA, but without T4 DNA ligase.

Table 2. Ligation mix.
Component Volume per reaction (μl) Final
10× T4 ligase buffer 2 2x
T4 ligase 0.7
ds-adapters (50 μM) 0.6 3 μM

4.3.2.Add 10 μl of the ligation mix to each sample (final volume 20 μl), as well as the 10 μl no ligase control to the control B.

4.3.3.Incubate at 16°C overnight (at least 16 h) in a thermocycler.

4.3.4.Inactivate the T4 DNA ligase by incubating 10 min at 65°C.

4.4.Amplify methylated DNA using HiDi DNA polymerase and Adr primer (Table 3 and Table 4). Include a “no DNA” control in which the DNA template is replaced with water (control C; Fig. 5A).

NOTE: Step 4 is subject to optimization in a sample-dependent manner (see critical steps below).

4.5.Verify each PCR reaction by loading 5 μl on a 1% agarose gel. Successful amplification of Dam-methylated DNA should yield a 400–1200 bp smear (Fig. 5C).

4.6.Purify the DNA with magnetic AMPure beads, following the AMPure XP 1.8× cleanup protocol detailed below.

4.6.1.For each PCR reaction, transfer 45 μl of PCR product into a 1.5 ml microcentrifuge tube.

Table 3. PCR-mix.
Component Volume per reaction (μl) Final
DNA (from ligation step) 20
10× HiDi PCR buffer 5
dNTP Mix 2.5 mM 4 0.2 mM
Nuclease-free water 18.75
Adr (50 μM) 1.25 1.25 μM
HiDi DNA polymerase 1
Table 4. PCR program (about 3 h).
Step Denaturation Annealing Extension
1 68°C for 10 min
2 94°C for 1 min 65°C for 5 min 68°C for 15 min
3 (4×) 94°C for 1 min 65°C for 1 min 68°C for 10 min

4.6.2.Briefly vortex the AMPure XP beads to resuspend them. Keep them on ice at all times.

4.6.3.Transfer 81 μl of bead suspension (corresponding to 1.8× the sample volume) into the tube containing the sample and pipette up and down 10 times to mix thoroughly. Incubate at room temperature for 5 min.

4.6.4.Transfer the tube to the magnetic stand and let it stand for at least 2 min for the beads to separate. From this step onwards, do not move the tubes from the stand unless explicitly mentioned.

4.6.5.Gently pipette out the cleared solution, always taking care not to disturb the beads.

4.6.6.Add 200 μl of freshly prepared 70% ethanol solution to wash the beads. Incubate for at least 30 s. Remove the supernatant. Repeat twice. (It is advisable to reach the bottom of the tube with the pipette tip and then dispense the solution).

4.6.7.Take the tubes out of the magnetic stand and let it stand for 30–60 s with the lid open so as to let the residual ethanol evaporate.

4.6.8.Elute by adding 30 μl of Milli-Q water or TE into the tube and resuspending the beads by pipetting (10 times).

4.6.9.Incubate at room temperature for 2 min and then place the tubes on the magnetic stand again.

4.6.10.Let the beads separate for 2 min and then slowly pipette out the eluted sample into a fresh microcentrifuge tube, recovering about 26–28 μl of purified DNA. Take care not to collect any bead.

4.7.Quantify DNA using Qubit. Concentration should be at least 20–50 ng/μl and the total amount > 0.08 pmol.

Table 5. End-repair master mix.
Component Volume per reaction (μl)
0.12 pmol amplicon 25
Ultra II End-prep buffer 3.5
Ultra II End-prep enzyme 1.5

5.1.2.Add 5 μl of the End-repair master mix to each 25 μl diluted amplicon PCR tube.

5.1.3.Mix gently by flicking the PCR tube, and quick spin to recover droplets.

5.1.4.Incubate at 20°C for 5 min in a thermocycler.

5.1.5.Inactivate the enzyme by incubating at 65°C for 5 min in a thermocycler.

5.1.6.Purify using magnetic beads:

5.1.7.(i) Add 1.8× volume (54 μl) of well resuspended AMPure XP beads. (ii) Incubate 10 min on the rotator mixer for binding. (iii) Spin down the sample and pellet on a magnetic rack. Incubate at least 5 min on the magnetic rack for the beads to separate. From this step onwards, do not move the tubes from the stand unless explicitly mentioned. (iv) Gently pipette out the cleared solution, always taking care not to disturb the beads. (v) Add 200 μl of freshly prepared 80% ethanol solution to wash the beads. Incubate for at least 30 s and pipette out the cleared solution. (It is advisable to reach the bottom of the tube with the pipette tip and then dispense the solution). (vi) Repeat twice making sure no ethanol droplet is left on the tube walls. (vii) Take the tubes out of the magnetic stand and let them dry for 5 min with the lid open so as to let the residual ethanol evaporate. (viii) Elute with 11 μl of Milli-Q water into each tube and resuspend the beads by pipetting (10 times). (ix) Incubate at room temperature for 2 min and then place the tubes on the magnetic stand again. (x) Let the beads separate for 2 min and then slowly pipette out the eluted sample into a new 0.2 ml PCR tube, recovering about 10.5 μl of purified DNA. Take care not to collect any bead with the eluate.

5.1.8.Use 1 μl to quantify DNA with Qubit. At least 50%–60% of input should be recovered.

Table 6. Barcoding mix.
Component Amount per reaction (μl)
End-repaired amplicons 9
Barcode (ONT) 1
NEB/Blunt TA master mix 10
Table 7. Adapter ligation mix.
Component Volume per reaction (μl)
Pooled barcode sample 65
Adapter Mix II (AMII, ONT) 5
NEBNext ligase buffer (5×) 20
Quick T4 DNA Ligase 10
Table 8. Library sequencing mix.
Component Volume per sample (μl)
SQB (ONT) 37.5
LB (ONT) 25.5
Pooled library 12

6.2.Starting sequencing run.

6.2.1.Turn on MinIT/computer/plug Minion.

6.2.2.Choose the flow cell type and check the “Available” box.

6.2.3.Make a hardware check of the Minion using the Configuration Test Cell. The flow cell must have >1000 active pores.

6.2.4.Click the “New Experiment” button at the bottom left of the GUI.

6.2.5.Select the running parameters for your experiment (FASTQ or FAST5).

6.2.6.Click “Start run”. We estimate that a good library should have at least 10 million reads after an overnight run.



We are grateful to Jaime Osuna-Luque for his participation in the earlier phase of the protocol development and for comments on the manuscript. Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). The study was supported by the Swiss National Science Foundation (IZCNZ0_174703/SBFI_C16.0013, BSSGI0_155764 and PP00P3_150681 to D.A.G.; ZCSZ0-174641/SBFI_C15.0076, PP00P3_159320 and 31003A_176226 to P.M.), the Novartis Foundation for Biomedical Research (to P.M. as well as to DAG) and a BMBS COST Action (BM1408).


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Supplementary information

File S1. Map of plasmid #418 in genbank format (.gb). [hsp-16.2p::loxP::mCherry::STOP::loxP::degron::gfp::dam::unc-54 3’UTR]
File S2. Map of plasmid #506 in genbank format (.gb). [hsp-16.2p::loxP::mCherry::STOP::loxP::degron::dam::rpb-6::unc-54 3’UTR]
Supplementary information of this article can be found online at