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1. Epitope Tagging of C. albicans strains
- Upload the gene of interest, along with its 1 kb upstream and downstream flanking sequences, from the Candida Genome Database to the primer design tool (see the Table of Materials). Design a guide RNA (gRNA) by highlighting 50 bp upstream and downstream from the stop codon, and click the gRNA selection tool on the right. Select Design and Analyze Guides. Use the Ca22 (Candida albicans SC5314 Assembly 22 (diploid)) genome and an NGG (SpCas9, 3' side) protospacer adjacent motif (PAM) for the guide parameters, and click Finish. Figure 1 describes the workflow for epitope tagging a C. albicans gene of interest with enhanced green fluorescent protein (eGFP).
- On the subsequent page, confirm the target region for gRNA design and press the green button. Sort gRNAs by the On-Target Score.
NOTE: The primer design tool computes on-target and off-target scores to quantify the specificity of the gRNAs. An ideal guide has an on-target score of >60, an off-target score of ~33, and overlaps the stop codon. This enables high gRNA specificity while ablating gRNA targeting after GFP integration. A gRNA with an off-target score of ~50 indicates allelic variation; thus, only one allele will be recognized by the gRNA.
- Add the sequences (5'-CGTAAACTATTTTTAATTTG-3') and (5'-GTTTTAGAGCTAGAAATAGC-3') to the 5' and 3' ends, respectively, of the 20 bp gRNA target sequence, creating a 60 bp primer/oligonucleotide. Alternatively, copy the 20 bp sequence to the gRNA calculator supplied by Nguyen et al.15. Order the 60 bp custom gRNA oligonucleotide.
- Amplify the "universal A fragment" with 100 mM AHO1096 (5'-GACGGCACGGCCACGCGTTTAAACCGCC-3') and 100 mM AHO1098 (5'-CAAATTAAAAATAGTTTACGCAAG-3') and the "unique B fragment" with the custom 60 bp gRNA oligonucleotide (100 mM) from step 1.1.2. and 100 mM AHO1097 (5'-CCCGCCAGGCGCTGGGGTTTAAACACCG-3') using pADH110 (plasmid repository ID# 90982) and pADH139 (plasmid repository ID# 90987), respectively, as template DNA. Use the PCR reaction and cycling conditions provided in Table 1.
NOTE: pADH139 is specific to strains that carry the heterologous Candida maltosa LEU2 marker. If using a strain with a single copy of the C. albicans LEU2 gene, substitute pADH119 (plasmid repository ID# 90985) in place of pADH139.
- Confirm successful amplification by checking 5 µL of the PCR on a 1% agarose gel. Look for ~1 kb A and B fragment amplicons.
- Mix 1 µL each of A and B fragments and stitch them together using the PCR reaction and cycling conditions provided in Table 2 to create a full-length C fragment.
- Add 0.5 µL of 100 mM AHO1237 (5'-AGGTGATGCTGAAGCTATTGAAG-3') and 0.5 µL of 100 mM AHO1453 (5'-ATTTTAGTAACAGCTTCGACAATCG-3') to each PCR reaction, mix well by pipetting, and complete the cycling conditions listed in Table 3.
NOTE: If using pADH119 in place of pADH139, substitute AHO1238 (5'-TGTATTTTGTTTTAAAATTTTAGTGACTGTTTC-3') in place of AHO1453.
- Confirm proper stitching and amplification of the C fragment by checking 5 µL of the PCR on a 1% agarose gel. Look for a ~2 kb amplicon. Store the C fragment at -20 °C until ready for use.
NOTE: If stitching and amplification results in multiple, nonspecific bands or smearing, perform a PCR cleanup of the A and B fragments and repeat from step 1.1.5.
- Add the entire CTG-optimized monomeric eGFP with linker sequence (RIPLING)16 (pCE1, plasmid repository ID# 174434) immediately upstream of the stop codon of the gene of interest using the primer design tool, creating a C-terminal translational fusion. Use this construct to design oligonucleotides for amplifying the donor DNA (dDNA) from pCE1.
- Design a forward oligonucleotide with 18-22 bp homology to the linker sequence and >50 bp homology to the 3' end of the open reading frame (ORF).
NOTE: The 18-22 bp homology creates an annealing temperature for amplification between 55 °C and 58 °C. If the full-length oligonucleotide forms primer dimers, adjust homology to the linker sequence/GFP or the genome accordingly.
- Create a reverse oligonucleotide with 18-22 bp homology to the 3' end of GFP and >50 bp homology to the downstream noncoding sequence of the ORF to be tagged.
- Order these oligonucleotides and amplify the dDNA using the provided touchdown PCR cycling conditions in Table 4.
- Design two sets of colony PCR (cPCR) oligonucleotides for confirming the integration of GFP by amplifying across the flanking integration sites. First, select the forward dDNA oligonucleotide using the primer design tool and click the Primer button on the right.
- Click Create Primers | Wizard | Tm Param and confirm that the algorithm is set to SantaLucia 1998. Click Use Selection to input the coordinates of the forward oligonucleotide designed in 1.2.1 as the target sequence.
- Set the optimal primer temperature to 55 °C and the maximum amplicon size to 900 bp, and click the Generate Primers button at the top right.
- Select the oligonucleotide pair with the lowest penalty score and confirm that the primers amplify across the 5' integration site. Ensure that the forward cPCR primer lies upstream of the forward dDNA primer sequence and the reverse cPCR primer fully within the eGFP tag or the linker sequence.
- Repeat steps 1.3-1.3.3. with the reverse dDNA oligonucleotide to create the second set of cPCR oligonucleotides that amplify across the 3' integration site. Ensure that the forward cPCR primer lies entirely within the eGFP tag or the linker sequence and the reverse cPCR primer downstream of the reverse dDNA primer sequence.
- Order these oligonucleotides.
- Digest 2,500 ng of pADH140, which contains Cas9 (plasmid repository ID# 90988), with restriction enzyme for each gene to be GFP-tagged. Set the total volume of each digestion at 15 µL; adjust the volume of water accordingly based on the pADH140 plasmid concentration. Use the digestion conditions specified in Table 5. Store the digested plasmid at -20 °C until ready for use.
NOTE: If transforming a strain with a single copy of the C. albicans LEU2 gene, instead of the heterologous C. maltosa LEU2 marker, substitute pADH137 (plasmid repository ID# 90986) in place of pADH140.
- Denature 12 µL of 10 mg/mL salmon sperm DNA for each gene that will be GFP-tagged at 99 °C for 10 min and rapidly cool to ≤4 °C. Store at -20 °C until ready for use.
- Streak a C. albicans LEU2 hemizygous nourseothricin-sensitive strain onto yeast peptone dextrose (YPD) plates and incubate at 30 °C for two days.
- Select a single colony and transfer it to 4 mL of liquid YPD. Incubate for 12-16 h at 30 °C with shaking at 250 rpm.
- Measure the optical density at 600 nm (OD600) of the overnight (12-16 h) culture in a spectrophotometer using a disposable cuvette (1 mL, 1 cm path length).
- Dilute the overnight culture into an Erlenmeyer flask to an OD600 of 0.1 in YPD. Account for 5 mL per reaction and include an additional 5 mL for checking the OD600 later.
NOTE: The volume of the culture depends on the number of transformation reactions.
- Incubate the diluted overnight culture in a shaking incubator at 30 °C with shaking at 250 rpm until it reaches an OD600 of 0.5-0.8.
- Centrifuge at 4,000 × g at room temperature for 5 min; remove and discard the supernatant.
- Resuspend the cell pellet in 1 mL of sterile water via gentle pipette mixing with filter tips and transfer to a sterile 1.5 mL microfuge tube.
- Pellet the cells by centrifuging at 4,000 × g at room temperature for 1 min; remove and discard the supernatant. Resuspend in 1 mL of sterile water and repeat for a total of two washes.
- Resuspend the pellet in 1/100th of the volume used in step 1.10. For example, if 15 mL was used, resuspend the pellet in 150 µL of sterile water.
- In a separate tube for each transformation reaction, mix 50 µL of C fragment, 50 µL of dDNA, 2,500 ng of restriction enzyme-digested pADH140, and 10 µL of denatured salmon sperm DNA.
- Add 50 µL of the cell slurry from step 1.14 and mix by pipetting.
- Make a stock of the plate mix (Supplementary File 1) for n + 1 transformations.
- Add 1 mL of the plate mix to the cell/DNA mixture and mix by inverting 5 times.
NOTE: Tap the bottoms of the tubes while inverting to dislodge any remaining liquid.
- Place the mixture in an incubator at 30 °C overnight (12-16 h) without shaking.
- Heat-shock the cells for 15 min at 44 °C in a water bath.
- Centrifuge the 1.5 mL microfuge tubes at 5,000 × g at room temperature for 2 min.
- Remove the PLATE mix by vacuum aspiration using sterile pipette tips, being careful to avoid disturbing the cell pellet.
- Resuspend the cell pellet in 1 mL of YPD, pellet by centrifugation at 4,000 × g at room temperature for 1 min, and remove and discard the supernatant. Repeat for a second wash, resuspend the cell pellet in 1 mL of YPD, and transfer the suspension to a 10 mL round-bottom, disposable culture tube containing an additional 1 mL of YPD (2 mL final volume). Recover the cells at 30 °C with shaking at 250 rpm for 5 h.
- Centrifuge the tubes at 4,000 × g at room temperature for 5 min; remove and discard the supernatant.
- Resuspend the cell pellet in 100 µL of sterile water and plate on YPD supplemented with 200 µg/mL nourseothricin (NAT200). Incubate at 30 °C for 2-3 days.
- Aliquot 100 µL of 20 mM NaOH into the wells of a 96-well PCR plate, with each well corresponding to an individual colony that grew on the NAT200 plates. Using a sterile toothpick or pipette tip, pick individual transformed colonies, patch them onto a new NAT200 plate, and swirl the remaining cells into a well with 20 mM NaOH. Repeat for the remaining colonies to create the cell lysate used as the DNA template for the cPCR reaction.
- Seal the PCR plate and incubate for 10 min at 99 °C in a thermocycler with a heated lid.
- Set up two cPCR reactions with the oligonucleotides designed in steps 1.3-1.3.5. Scale up the number of reactions as needed. Perform the PCR reaction with the cell lysate prepared in step 1.26 following cycling conditions and PCR reaction mixtures from Table 6. Run 10-20 µL from each well on a 1% agarose gel. Look for colonies with amplification of the two cPCR primer sets indicating properly incorporated GFP dDNA.
- Restreak colonies that incorporated GFP on synthetic complete (SC) media lacking leucine. Incubate in a 30 °C incubator for 2-3 days. Pick individual colonies and patch onto YPD and YPD supplemented with 400 μg/mL nourseothricin (NAT400) plates. Identify colonies that fail to grow on NAT400 plates after 24 h as those that have successfully lost the CRISPR components.
- Confirm that the GFP tag is retained by repeating steps 1.25-1.28 using cells from the YPD patch plate. If the correct bands are present, inoculate into 4 mL of YPD and grow overnight (12-16 h), as described in step 1.7.
- Mix the overnight culture of the new GFP-tagged strain with filter-sterilized 50% glycerol in a 1:1 ratio in a sterile cryotube. Store at -80 °C and restreak onto YPD plates as needed.
NOTE: It is recommended to validate the GFP-tagged strains by confirming nuclear localization of the tagged TF via fluorescent microscopy and confirming a wild-type phenotype in an appropriate phenotypic assay.
2. Sample preparation of biofilm cultures
- Streak C. albicans GFP-tagged strain(s) onto YPD agar plates and incubate at 30 °C for 2-3 days. Using a single isolated colony from the agar plate, inoculate into 4 mL of YPD liquid medium. Incubate at 30 °C with shaking overnight (12-16 h). Determine the OD600 of the overnight culture(s).
NOTE: It is recommended to use three biological replicates per sample for the CUT&RUN experiments.
- Inoculate a sterile 12-well untreated cell culture plate with the overnight culture to a final OD600 of 0.5 (equivalent to 2 × 107 cells/mL) in Roswell Park Memorial Institute (RPMI)-1640 medium to a final volume of 2 mL. Incubate for 90 min at 37 °C in a microplate incubator with shaking at 250 rpm.
NOTE: It is recommended to use one 12-well cell culture plate per strain with one well uninoculated as a medium-alone contamination control. This protocol has been successfully applied using as little as 1/10th of one 12-well cell culture plate well (or as few as 5 million cells). Using a higher number of cells increases total DNA yields, which typically results in high-quality sequencing libraries.
- Remove unadhered cells by aspiration using sterile pipette tips attached via flexible plastic tubing to a vacuum trap apparatus. Wash the adhered cells once with 2 mL of sterile 1x phosphate-buffered saline (PBS). Add 2 mL of fresh RPMI-1640 medium to the wells and incubate for 24 h at 37 °C with shaking at 250 rpm.
NOTE: Change pipette tips between wells of different strains and/or conditions. Do not scrape the bottom of the well with the tip while aspirating.
- At the end of the 24 h incubation, collect and pool the liquid and biofilm material from each of the 11 inoculated wells into a single, sterile 50 mL conical tube. Repeat as necessary with independent pools if processing more than one strain or growth condition concurrently.
NOTE: Scrape the bottoms and edges of each well with a pipette filter tip to dislodge cells that remain adhered to the surface. Use the pipette to homogenize the biofilms.
- Pellet samples by centrifuging at 4,000 × g at room temperature for 5 min. Decant as much of the supernatant as possible, taking care to minimize disruption of the pellet. Snap-freeze the pellet in liquid nitrogen and store at -80 °C immediately after collection or continue directly to step 4 (isolation of nuclei).
3. Sample preparation of planktonic cultures
- Streak C. albicans GFP-tagged strain(s) onto YPD agar plates and incubate at 30 °C for 2-3 days. Using a single isolated colony from the agar plate, inoculate into 4 mL of YPD liquid medium. Incubate at 30 °C with shaking overnight (12-16 h). Determine the OD600 of the overnight culture(s).
- Back-dilute overnight cultures to OD600 of 0.1 in 50 mL of RPMI-1640 liquid medium and incubate at 30 °C with shaking at 225 rpm for 2-5 h until OD600 is between 0.5 and 0.8.
NOTE: Cells should go through at least two doublings before being harvested. Conditions used for planktonic cultures can be adjusted as needed.
- Pellet the samples by centrifuging at 4,000 × g at room temperature for 5 min. Decant as much of the supernatant as possible, taking care to minimize disruption of the pellet. Snap-freeze the pellet in liquid nitrogen and store at -80 °C immediately after collection or continue directly to step 4 (isolation of nuclei).
4. Isolation of nuclei
NOTE: On the day of the experiment, prepare fresh Ficoll Buffer, add 2-mercaptoethanol and protease inhibitor to aliquot(s) of the Resuspension Buffer, and add protease inhibitor to aliquot(s) of the SPC Buffer (see Supplementary File 1). To resuspend the pellets, gently pipette using either 200 µL or 1 mL pipette tips to avoid damaging the cells or nuclei. Before beginning the nuclei isolation, turn on the heat block to preheat it to 30 °C. All pipette tips and tubes for the remainder of this protocol should be certified DNA/RNA and DNase/RNase-free, and the use of filter tips is recommended for all subsequent pipetting steps.
- Resuspend pellet(s) in 1 mL of room-temperature Resuspension Buffer and transfer to a sterile 1.5 mL microfuge tube. Pellet at 2,000 × g at room temperature for 2 min in a table-top centrifuge and remove the supernatant.
NOTE: Remove the supernatant using either 200 µL or 1 mL pipette tip, taking care to minimize disruption of the pellets.
- Resuspend the pellet(s) in 200 µL of room-temperature Resuspension Buffer. From the resuspended pellet, transfer a 5 µL aliquot into a new PCR tube and store it at 4 °C for use later.
NOTE: This aliquot will be used as a control during a subsequent quality control step to evaluate the quality of the isolated nuclei.
- Pellet at 2,000 × g for 2 min and remove and discard the supernatant using a pipette. Repeat the wash step twice using 200 µL of Resuspension Buffer.
- Centrifuge at 2,000 × g at room temperature for 2 min and remove the supernatant. Add 300 µL of Resuspension Buffer and 10 µL of lyticase solution (50 mg/mL, see the Table of Materials). Incubate for 30 min at 30 °C in a heat block.
NOTE: Alternatively, a water bath heated to 30 °C can also be used instead of a heat block. The spheroplasting conditions used here have been optimized to be effective for both yeast and hyphal cells of C. albicans. It is recommended to optimize the spheroplasting conditions when applying this protocol to C. albicans cells with mutations that impact cell wall integrity or other cellular morphologies. During this 30 min incubation step, the user has the option to complete step 5 (Concanavalin A Bead Activation) ahead of time to save time.
- CRITICAL STEP: After the 30 min incubation step, transfer a 5 µL aliquot into a new PCR tube. To the 5 µL of isolated nuclei and the 5 µL aliquot of intact cells stored at 4 °C from step 4.2, add 1 µL calcofluor white (a fluorescent cell wall dye) and 1 µL of SYTO 13 (a nucleic acid stain). Incubate at 30 °C in the dark for 30 min.
- Visually inspect the integrity and purity of the isolated nuclei using a fluorescence microscope. Look for isolated nuclei that show prominently stained intact nuclei (using a 488-509 nm excitation filter) and ensure that there is no cell wall staining by the calcofluor white dye (using a 390-420 nm excitation filter). In the intact control cells, look for prominent cell wall staining by the calcofluor white dye (using a 390-420 nm excitation filter) and stained intact nuclei (using a 488-509 nm excitation filter).
- Centrifuge at 2,000 × g at 4 °C for 5 min and remove the supernatant. Resuspend the pellet in 500 µL of ice-cold Resuspension Buffer using 1 mL filter tips by pipetting gently up and down 5 times. Centrifuge at 2,000 × g at 4 °C for 5 min, and remove the supernatant using a 1 mL pipette. Resuspend the pellet with 1 mL of freshly made ice-cold Ficoll Buffer.
NOTE: Keep the samples and buffers on ice from this point forward.
- Centrifuge the samples at 5,000 × g at 4 °C for 10 min and remove the supernatant. Resuspend the pellet in 500 µL of ice-cold SPC Buffer.
NOTE: From this point onward, handle the nuclei extremely gently to avoid damaging them.
- Centrifuge the samples at 5,000 × g at 4 °C for 10 min and remove as much of the supernatant as possible without disrupting the pellet. Place the tubes containing the pelleted nuclei on ice and proceed to step 5. If step 5 was already completed ahead of time in step 4.4, proceed to step 6 or snap-freeze the pellets in liquid nitrogen and store them at -80 °C immediately after collection.
5. Concanavalin A bead activation
NOTE: This is a critical step. From this point forward, users have the option to continue with the protocol using a commercially available CUT&RUN kit or source key components individually and prepare buffers in-house. If using the commercial kit, all buffers and reagents used below are included in the kit unless otherwise noted. Individual catalog numbers for sourcing reagents independently are also provided in the Table of Materials. Chill all buffers on ice before use. Once step 5 is completed, it is recommended to proceed to step 6 immediately. Avoid multiple freeze-thawing of isolated nuclei as it is known to increase DNA damage and could lead to poor quality results.
- Gently resuspend the concanavalin A (ConA) beads using a pipette. Transfer 22 µL of ConA bead suspension per sample to be processed in a single 1.5 mL microfuge tube. Place the tube on a magnetic rack until the bead slurry is clear; remove and discard the supernatant using a pipette.
NOTE: When performing CUT&RUN for a total of 10 samples, for example, transfer 220 µL of the ConA bead suspension to a 1.5 mL microfuge tube.
- Remove the tube containing the ConA beads from the magnetic rack and immediately add 200 µL of ice-cold Bead Activation Buffer and gently mix using a pipette. Place the tube on the magnetic rack until the bead slurry is clear; remove and discard the supernatant using a pipette. Repeat this step for a total of two washes.
- Resuspend the beads in 22 µL of ice-cold Bead Activation Buffer per sample of nuclei to be processed. Keep the beads on ice until needed.
NOTE: The throughput of the subsequent steps is dependent on the number and capacity of magnetic tube racks available. Processing 32 samples in two 16-well tube racks is a manageable number for most users of this protocol. However, higher throughput is possible for more experienced users, or if robotic liquid handling systems are available.
6. Binding nuclei to activated beads
NOTE: Chill all buffers on ice before use. All buffers supplemented with protease inhibitors should be prepared fresh on the day of the experiment. It is recommended to use 0.2 mL strip tubes in the subsequent steps.
- Resuspend the pelleted nuclei from step 4 in 100 µL of ice-cold SPC Buffer and transfer to a new 8-tube 0.2 mL strip. Add 20 µL of the activated beads to each sample and gently pipette to mix. Incubate at room temperature for 10 min without agitation.
- Place the tubes on the magnetic rack until the slurry is clear; remove and discard the supernatant using a pipette. Remove the tubes from the magnetic rack, and add 200 µL of ice-cold Wash Buffer to each sample. Resuspend the beads by gently pipetting up and down 5 times. Transfer 100 µL aliquots from each sample into a new 8-tube 0.2 mL strip.
- CRITICAL STEP: Divide each CUT&RUN sample into two separate aliquots. Use one of the aliquots for the negative control antibody (e.g., IgG negative control antibody) and the other for the target antibody against the protein of interest (e.g., anti-GFP antibody).
NOTE: Both samples are required for the computational pipeline to accurately identify enrichment signals specific to the TF of interest. An additional control using anti-GFP antibodies with an untagged strain can also be performed. This control has shown results comparable to the use of IgG antibodies in a GFP-tagged strain. Therefore, for simplicity, it is recommended to use the standard IgG control for all experiments.
7. Primary antibody binding
NOTE: pAG-MNase fusion protein binds well to rabbit, goat, donkey, guinea pig, and mouse IgG antibodies17. Generally, most commercial ChIP-seq-certified commercial antibodies are compatible with CUT&RUN procedures. The amount of primary antibody used depends on the efficiency of the antibody, and titration of the antibody (e.g., 1:50, 1:100, 1:200, and 1:400 final dilution) may be necessary if the antibody of interest has not been previously tested in ChIP or CUT&RUN experiments. Chill all buffers on ice prior to use. All buffers used for antibody binding steps should be prepared fresh on the day of the experiment.
- Place the tubes on a magnetic rack and wait until the slurry is completely clear; remove and discard the supernatant using a pipette. Add 50 µL of the Antibody Buffer and gently mix by pipetting.
- Add 3 µL of the anti-GFP polyclonal antibody (or 0.5 µg if using an untested antibody). Incubate the tubes on a nutating mixer at 4 °C for 2 h.
NOTE: Some CUT&RUN protocols report increased yield by adding a secondary antibody prior to pAG-MNase addition14; however, no significant improvement was observed using this added step, and thus, it is not included in this protocol.
- Briefly centrifuge the tubes at 100 × g at room temperature for 5 s, place the tubes on a magnetic rack, and once the slurry is clear, remove and discard the supernatant using a pipette. While the tubes containing the beads are still on the magnetic rack, add 200 µL of ice-cold Cell Permeabilization Buffer directly onto the beads. Remove and discard the supernatant using a pipette. Repeat for a total of two washes with the ice-cold Cell Permeabilization Buffer.
- Add 50 µL of ice-cold Cell Permeabilization Buffer to each tube and gently mix by pipetting.
NOTE: Beads are often aggregated at this point but can easily be dispersed by mixing gently using a 200 µL pipette.
8. Binding of pAG-MNase to antibody
- Add 2.5 µL of the pAG-Mnase (20x stock) to each sample and gently mix by pipetting. Place the samples (slightly elevated at ~45° angle) on a nutator at 4 °C. Turn on the nutator and incubate the samples for 1 h.
- Briefly centrifuge the strip tubes at 100 × g at room temperature for 5 s, place the tubes on a magnetic rack and, once the slurry is clear, remove and discard the supernatant using a pipette.
NOTE: This step is critical. Carryover antibody remaining in the cap or sides of the tubes after this step will significantly increase the amount of background signal.
- While the tubes containing the beads are still on the magnetic rack, add 200 µL of ice-cold Cell Permeabilization Buffer, allow the slurry to clear, and remove and discard the supernatant using a pipette. Repeat this step for a total of two washes with the Cell Permeabilization Buffer.
- Add 100 µL of the ice-cold Cell Permeabilization Buffer to the samples and gently pipette up and down 5 times.
9. Targeted chromatin digestion and release
- Incubate the tubes containing the sample(s) in a wet ice bath for 5 min. Add 3 µL of 100 mM CaCl2 into each sample using a multichannel pipette. Gently pipette up and down 5 times, immediately return the tubes to the wet ice bath, and incubate for 30 min.
- Add 66 µL of the Stop Buffer to each sample and gently vortex to mix. Incubate samples for 10 min at 37 °C in a dry bath.
NOTE: It is recommended to add 1.5 pg of heterologous E. coli spike-in DNA per sample in the Stop Buffer. The addition of 1.5 pg of E. coli spike-in DNA results in 1,000-10,000 mapped spike-in reads for 1-10 million mapped experimental reads14. The spike-in DNA is used to calibrate the sequencing depth and is especially important for comparing samples in a series. The addition of spike-in E. coli is highly recommended but not essential. The commercial CUT&RUN kit includes E. coli spike-in DNA, but it can also be purchased separately.
- Place the tubes on the magnetic rack and transfer 160 µL of the supernatant into a 1.5 mL microfuge tube. Transfer 80 µL of the sample into a new 2 mL microfuge tube and store at -20 °C in the event that a backup sample is needed. Proceed to step 10 with the 80 µL sample.
10. Cleanup of collected DNA samples
NOTE: Incubate DNA Purification Beads at room temperature for 30 min before use. Prechill 100% isopropanol on ice. When mixing the samples, pipette up and down 10 times.
- Vortex DNA Purification Beads to homogenize the bead suspension. Add 50 µL (~0.6x sample volume) of the resuspended beads to each sample. Pipette-mix and incubate the samples on a nutator for 5 min at room temperature.
NOTE: The ratio of DNA purification beads to sample used is critical. Using 0.6x volume of DNA Purification Bead solution relative to the sample allows the magnetic beads to bind to large DNA fragments released from damaged nuclei. CUT&RUN-enriched DNA fragments are much smaller than these large DNA fragments and are thus retained in the supernatant at this step.
- Place the tubes on a magnetic rack and transfer 130 µL of the supernatant containing the DNA to a 0.2 mL 8-tube strip. Add an additional 30 µL of DNA Purification Beads to the sample(s) (the total volume is 160 µL).
- Add 170 µL (~1x sample volume) of ice-cold 100% isopropanol, mix well by pipetting up and down 10 times, and incubate on ice for 10 min.
NOTE: It is critical that 100% ice-cold isopropanol is used for this step for the DNA purification beads to efficiently capture the CUT&RUN-enriched small fragments.
- Place the tubes on the magnetic rack, and once the slurry has cleared, carefully remove and discard the supernatant using a pipette.
- While the tubes are on the magnetic rack, add 200 µL of freshly prepared, room-temperature 80% ethanol to the tubes and incubate at room temperature for 30 s. Carefully remove and discard the supernatant using a pipette. Repeat this step for a total of two washes with 80% ethanol.
- Quickly spin the tubes at 100 × g, place the tubes back on the magnetic rack, and remove any residual ethanol using a pipette after the slurry has cleared. Air-dry the beads for 5 min while the tubes remain on the magnetic rack with the lid open.
NOTE: Do not exceed 5 min of drying time as this can significantly reduce the final DNA yield.
- Remove the tubes from the magnetic rack and elute the DNA from the beads by adding 17 µL of 0.1x Tris-EDTA (TE) at pH 8. Mix well and incubate the tubes for 5 min at room temperature.
- Place the tubes on the magnetic rack until the slurry becomes clear. Once the slurry has cleared, carefully transfer 15 µL of the supernatant to a sterile 0.2 mL PCR tube.
- Measure the concentration of the collected DNA using a fluorometer following the manufacturer's protocol.
NOTE: Typically, the concentration of the collected DNA is ~1 ng/µL. Sometimes, the concentration of the collected DNA is too low to quantify using a fluorometer. This is not an indicator of a failed experiment. Proceed with the library preparation regardless of the concentration of the collected DNA.
- Proceed to step 11 or store the samples at -20 °C until ready.
11. Library preparation for sequencing
NOTE: The following steps use a commercially available library prep kit. When performing steps using the Ligation Master Mix, minimize touching the tubes and always keep them on ice.
- Using 0.1x TE at pH 8, bring up the total volume of the CUT&RUN DNA to 50 µL. Make a master mix of 3 µL of End Prep Enzyme Mix and 7 µL of End Prep Reaction Buffer per sample. Add 10 µL of master mix to the CUT&RUN DNA and mix thoroughly by pipetting up and down 5 times.
- Perform a quick spin at 100 × g to collect all liquid from the sides of the tube. Place the tubes in a thermocycler with the heated lid set to ≥75 °C and run the cycling conditions in Table 7.
NOTE: Depending on the starting input DNA concentrations collected from step 10, follow the required adapter dilution from Table 8.
- Add 2.5 µL of Adapter per sample and mix thoroughly by pipetting up and down 10 times.
NOTE: It is critical that the adapter is added to the sample and mixed thoroughly before the ligation master mix is added.
- Make a master mix of 30 µL of Ligation Master Mix and 1 µL of Ligation Enhancer. Add 31 µL of the master mix to the sample(s). Mix thoroughly by pipetting up and down 10 times.
- Incubate at 20 °C for 15 min in a thermocycler with the heated lid off.
NOTE: It is critical that samples be kept on ice and transferred to the thermocycler only after the thermocycler has reached 20 °C.
- Perform a quick spin at 100 × g to collect all liquid from the sides of the tube, add 3 µL of Uracil Excision Enzyme, and incubate the tubes in the thermocycler at 37 °C for 15 min with the heated lid set to ≥47 °C.
NOTE: This is a safe stopping point; store the samples at -20 °C or continue directly to step 11.7. If continuing directly to step 11.7, incubate the DNA Purification Beads at room temperature for 30 min before use.
- Add 154.4 µL (~1.6x sample volume) of the DNA Purification Beads to the Adapter Ligation reaction from step 11.6. Pipette-mix and incubate the samples for 5 min at room temperature.
- Place the tubes on the magnetic rack and once the slurry has cleared, carefully remove and discard the supernatant using a pipette.
- Add 200 µL of freshly prepared, room-temperature 80% ethanol to the tubes and incubate at room temperature for 30 s. Carefully remove and discard the supernatant using a pipette and repeat this step for a total of two washes with 80% ethanol.
- Spin the tubes briefly at 100 × g. Place the tubes back on the magnetic rack and remove any residual ethanol using a pipette. Air dry the beads for 5 min while the tubes remain on the magnetic rack with the lid open.
NOTE: Do not exceed 5 min of drying time as this can significantly reduce the final DNA yield.
- Remove the tubes from the magnetic rack and elute the DNA from the beads by adding 17 µL of 0.1x TE at pH 8. Mix well and incubate for 5 min at room temperature.
- Place the tubes on the magnetic rack until the slurry becomes clear. Once the slurry has cleared, carefully transfer 15 µL of the supernatant to a sterile 0.2 mL PCR tube.
- Make a master mix of 25 µL of DNA Polymerase Master Mix and 5 µL of Universal Forward Library Amplification Primer (10 µM) per sample.
NOTE: Prepare one extra sample of master mix to account for pipetting losses.
- Add 30 µL of the master mix to the 15 µL of Adapter-ligated DNA sample. Add 5 µL of Reverse Uniquely Indexed Library Amplification Primer (10 µM) to each sample to bring the final volume to a total of 50 µL. Mix thoroughly by pipetting up and down 10 times. Perform the PCR cycling conditions in Table 9.
NOTE: Incubate the DNA Purification Beads at room temperature for 30 min before use.
- Vortex the DNA Purification Beads to resuspend. Add 35 µL (~0.7x sample volume) of the resuspended beads to the PCR-amplified DNA samples. Mix and incubate the samples on a nutator for 5 min at room temperature.
- Place the tubes on the magnetic rack and once the slurry is clear, transfer the supernatant containing the DNA to a new 0.2 mL 8-well PCR strip tube.
- Add 119 µL (~1.4x sample volume) of beads to the sample, and mix by pipetting up and down 5 times. Incubate the samples on a nutator for 5 min at room temperature.
- Place the tubes on the magnetic rack and once the slurry has cleared, carefully remove and discard the supernatant using a pipette.
- Add 200 µL of freshly prepared, room-temperature 80% ethanol to the tubes and incubate at room temperature for 30 s. Carefully remove and discard the supernatant using a pipette; repeat the step for a total of two washes with 80% ethanol.
- Spin the tubes briefly at 100 × g. Place the tubes back on the magnetic rack and remove any residual ethanol using a pipette. Air-dry the beads for 5 min while the tubes remain on the magnetic rack with the lid open.
NOTE: Do not exceed 5 min of drying time as this can significantly reduce the final DNA yield.
- Remove the tubes from the magnetic rack and elute the DNA from the beads by adding 14 µL of 0.1x TE at pH 8. Mix well and incubate for 5 min at room temperature.
- Place the tubes on the magnetic rack until the slurry becomes clear. Once the slurry has cleared, carefully transfer 13 µL of the supernatant to a sterile 0.2 mL PCR tube.
- Prepare fresh 1x Tris-borate-EDTA (TBE) and insert premade, commercial 10% acrylamide TBE gel into the gel electrophoresis apparatus filled with 1x TBE.
- In the first well, add 2 µL of Low-range DNA ladder. Mix 3 µL of 6x loading dye with 13 µL of the sample previously collected from step 11.22. Carefully add 15 µL into each well of the gel. Run the gel for 90 min at 70 V.
NOTE: It is recommended to leave one well in the gel empty between each sample, as this reduces the likelihood of sample cross contamination. Experienced users may find it appropriate to use all wells while carefully avoiding cross contamination, particularly when processing a large number of samples.
- Remove the gel cast from the gel box. Open the gel cast per the manufacturer's instructions, gently remove the gel from the gel cast, and place it inside a gel holding tray containing 100 mL of 1x TBE.
NOTE: Make sure to gently remove the gel from the gel cast to avoid ripping the gel as it is thin and fragile. It is critical to prewet gloves and the gel with 1x TBE whenever handling the gel. The gel holding tray should be slightly larger than the size of the gel (approximately 0.5" on each side). The plastic lids provided with 96-well PCR-tube storage boxes are convenient holding trays for standard mini gel sizes.
- Add 10 µL of the nucleic acid gel stain to the tray and gently swirl. Cover with foil to protect from light and incubate statically at room temperature for 10 min.
- Rinse the gel twice with 100 mL of deionized tap water. Image the gel under blue light illumination using an amber filter cover (Figure 2).
NOTE: Successful libraries show a smear between 100 and 500 bp. There will also be a prominent ~125 adapter dimer band. The presence of adapter dimers is not an indicator of poor library quality. This amount of adapter dimers is unavoidable for CUT&RUN experiments performed on low-abundance TFs and is a consequence of the low amount of input material used to prepare these libraries. Do not use ultraviolet light, which can damage the DNA.
- As shown in Figure 2, for each library, cut the gel slightly above the ~125 bp prominent adapter dimer band (making sure to avoid touching the adapter dimer band) and below the 400 bp ladder mark.
NOTE: It is critical to avoid the ~125 adapter dimer band. Even tiny amounts of adapter dimers will significantly reduce the library quality.
- Puncture the bottom of a 0.65 mL tube using a 22 G needle and place the punctured tube inside a sterile 2 mL microfuge tube. Transfer the gel slice to the punctured tube inside the 2 mL microfuge tube.
- Centrifuge the 2 mL microfuge tube containing the 0.65 mL punctured tube and sample at 10,000 × g at room temperature for 2 min to collect the gel slurry inside the 2 mL microfuge tube.
NOTE: The punctured tube should now be empty and can be discarded. If the punctured tube still has any gel remaining inside, place the punctured tube back inside the 2 mL microfuge tube and centrifuge again at 10,000 × g at room temperature for an additional 2 min.
- To the gel slurry inside the 2 mL microfuge tube, add 300 µL of ice-cold gel elution Buffer and mix on a nutator at room temperature for a minimum of 3 h or overnight (12-16 h).
- Transfer all liquid and gel slurry to a 0.22 µm filter column. Centrifuge at 10,000 × g at room temperature for 1 min; the collected volume should be ~300 µL.
- Add 450 µL (~1.5x sample volume) of DNA Purification Beads, incubate at room temperature for 5 min on a nutator, and then place the sample on the magnetic rack until the slurry is clear.
NOTE: Incubate the DNA Purification Beads at room temperature for 30 min before use.
- Remove and discard 500 µL of the supernatant, making sure not to disrupt the beads.
- Remove the sample from the magnetic rack and mix the beads by pipetting up and down 5 times. Transfer 200 µL of the sample into a new PCR strip tube.
- Place the strip tube on the magnetic rack, and once the slurry has cleared, carefully remove and discard the supernatant using a pipette.
- Add 200 µL of freshly prepared, room-temperature 80% ethanol to the tubes and incubate at room temperature for 30 s. Carefully remove and discard the supernatant using a pipette; repeat this step for a total of two washes with 80% ethanol.
- Spin the tubes briefly at 100 × g. Place the tubes back on the magnetic rack and remove any residual ethanol using a pipette. Air-dry the beads for up to 5 min while the tubes remain on the magnetic rack with the lid open.
NOTE: Do not exceed 5 min of drying time as this can significantly reduce the final DNA yield.
- Remove the tubes from the magnetic rack and elute the DNA from the beads by adding 17 µL of 0.1x TE at pH 8. Mix well and incubate the tubes for 5 min at room temperature.
- Place the tubes on the magnetic rack until the slurry becomes clear. Once the slurry has cleared, carefully transfer 15 µL of the supernatant to a sterile 0.2 mL PCR tube.
- Measure the final library quantity using the fluorometer; use this final library for sequencing.
NOTE: Up to 48 libraries can be pooled and sequenced together in a single lane using a sequencing platform that provides at least 300 million 40 bp or longer paired-end reads.
12. CUT&RUN sequence analysis
NOTE: This section presents the computational protocol used to analyze the CUT&RUN sequence data. The protocol begins with setting up the computational virtual environment and walks users through executing the commands on their local machine. This protocol will work on all computational resources, such as local machines, virtual cloud servers, and high-performance computing clusters. All CUT&RUN data presented in this paper can be accessed at NCBI GEO under accession number GSE193803.
- Download the source code for the CUT&RUN analysis from https://github.com/akshayparopkari/cut_run_analysis.
NOTE: The workflow will work best on a MacOS or Linux OS system. Windows users can run the workflow using GitBash (see the Table of Materials).
- Directly download the code from the GitHub page by clicking on the green Code button | Download ZIP option. Unzip the folder to a relevant location on the local machine.
- Install Conda environment (see the Table of Materials) and run only once.
NOTE: This workflow uses the Conda command line tool environment to install all required software and tools.
- Once Conda is installed (run only once), create a virtual environment using the Supplementary File 2 provided with the following command:
conda create --name <env> --file Supplementary_File_2.txt
- Activate the virtual environment every time this workflow is to be executed using:
conda activate <env>
- Organize the input raw FASTQ files into a single folder, ideally one folder per CUT&RUN experiment.
13. Generation of the genome file for alignment
- Generate the genome file for alignment (run only once for each genome file). Create a folder to save all C. albicans genome files, such as:
mkdir ca_genome_files
14. Downloading C. albicans genome assembly 21
- Download C. albicans genome assembly 21 from the Candida Genome Database using either wget or curl tools (see the Table of Materials)18.
NOTE: C. albicans assembly 21 was used here to compare CUT&RUN results with previously published ChIP-chip results, which were aligned to Assembly 21. Users can download other assembly versions and run similar commands to generate relevant genome files for their alignment needs.
15. Generate a Bowtie 2 index database (database name: ca21)
- Use the following:
bowtie2-build C_albicans_SC5314_A21_current_chromosomes.fasta.gz ca21
bowtie2-inspect -s ca21
16. Run the CUT&RUN analysis pipeline
- Read the help section to become familiar with the parameters of the pipeline.
bash cut_n_run_pipeline.sh -h
17. Execute the cut_n_run_pipeline.sh file with relevant parameters
- Execute the script:
bash cut_n_run_pipeline.sh /path/to/input/folder 4 y y y y y > /path/to/output.log 2>&1
NOTE: The relevant parameters with detailed descriptions on lines 19-36 of the code are described on the GitHub page https://github.com/akshayparopkari/cut_run_analysis/blob/main/cut_n_run_pipeline.sh.
18. Organize output files
- Merge significant peaks from all replicates called by MACS2 located in /path/to/input/folder/peakcalling/macs2 using the BedTools merge function19.
cat /path/to/input/folder/peakcalling/macs2/all_replicate_files sort -k1,1 -k2,2n | mergeBed -c 4,5,6,7,8,9 -o last,mean,first,mean,mean,mean > /path/to/merged_output.bed
NOTE: For additional information and best practices on assessing overlapping peaks in replicate samples, please refer to Landt et al.20 and Boyd et al.21.
19. Remove matches to blocklisted genomic regions using the BedTools subtract function
- Use the following: subtractBed -a /path/to/merged_output.bed -b /path/to/Supplementary_File_3.bed -A > /path/to/merged_output_no_blocklist_hits.bed
NOTE: The blocklisted regions in the C. albicans genome are provided as a .bed file in Supplementary File 3. The list consists primarily of highly repetitive sequence elements and regions such as telomeric repeats and centromeres that commonly yield false-positive results in C. albicans CUT&RUN, ChIP-seq, and ChIP-chip datasets. Hence, it is recommended to remove the blocklisted regions. However, for certain protein targets, it may be inappropriate or undesirable to exclude these loci. Users can skip this step to retain signals contained within these blocklisted regions or create their own blocklisted regions.
20. Merge BigWig files from replicates using the UCSC bigWigMerge function22
- Use the following: bigWigMerge /path/to/input/folder/bigwig/all_final_bw_files /path/to/input/folder/bigwig/ all_final_bdg_file
- Convert the BedGraph output from bigWigMerge to BigWig using the UCSC bedGraphToBigWig function.
bedGraphToBigWig /path/to/input/folder/bigwig/ all_final_bdg_file /path/to/input/folder/bigwig/ all_final_bw_file