Analyzing Telomeric Protein-DNA Interactions Using Single-Molecule Magnetic Tweezers

Summary

This protocol demonstrates using single-molecule magnetic tweezers to study interactions between telomeric DNA-binding proteins (Telomere Repeat-binding Factor 1 [TRF1] and TRF2) and long telomeres extracted from human cells. It describes the preparatory steps for telomeres and telomeric repeat-binding factors, the execution of single-molecule experiments, and the data collection and analysis methods.

Abstract

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper function depends on tightly regulated processes of replication, elongation, and damage response. The shelterin complex, especially Telomere Repeat-binding Factor 1 (TRF1) and TRF2, plays a pivotal role in telomere protection and has emerged as a potential anti-cancer target for drug discovery. These proteins bind to the repetitive telomeric DNA motif TTAGGG, facilitating the formation of protective structures and recruitment of other telomeric proteins. Structural methods and advanced imaging techniques have provided insights into telomeric protein-DNA interactions, but probing the dynamic processes requires single-molecule approaches. Tools like magnetic tweezers, optical tweezers, and atomic force microscopy (AFM) have been employed to study telomeric protein-DNA interactions, revealing important details such as TRF2-dependent DNA distortion and telomerase catalysis. However, the preparation of single-molecule constructs with telomeric repetitive motifs continues to be a challenging task, potentially limiting the breadth of studies utilizing single-molecule mechanical methods. To address this, we developed a method to study interactions using full-length human telomeric DNA with magnetic tweezers. This protocol describes how to express and purify TRF2, prepare telomeric DNA, set up single-molecule mechanical assays, and analyze data. This detailed guide will benefit researchers in telomere biology and telomere-targeted drug discovery.

Introduction

Telomeres are protective structures at the ends of chromosomes^1,2,3. Telomere erosion during cell division leads to cell senescence and aging, while abnormal elongation of telomeres contributes to cancer^4,5. For telomeres to function properly, their replication, elongation, and damage responses must be highly regulated^6,7,8. Shelterin, composed of six subunits, plays a central role in telomere protection^9,10,11. A deeper understanding of telomeres will provide valuable insights into telomere biology.

TRF1 and TRF2, core subunits of shelterin, are telomeric binding proteins^12,13. Both TRF1 and TRF2 bind to the repetitive DNA motif TTAGGG in telomeres via their Myb domains¹⁴. They form dimers through their shared TRFH domains, which allow them to encircle telomeric double-stranded DNA and to recruit telomeric proteins^{15,16,17,18,19}. TRF2 is particularly important for the formation of telomeric D-loops and T-loops^20,21. Due to their crucial roles in telomere protection, TRF1 and TRF2 have emerged as potential anti-cancer drug targets^22,23,24,25.

Significant efforts have been made to investigate the protein-DNA interactions at telomeres. Biochemical methods such as Electrophoretic Mobility Shift Assay (EMSA) and Surface Plasmon Resonance (SPR) have been used to examine binding affinities^20,26. Numerous structures of telomeric binding proteins complexed with DNA have been elucidated using cryo-electron microscopy (cryo-EM), X-ray crystallography, and nuclear magnetic resonance (NMR)^27,28,29. Super-resolution imaging techniques like stochastic optical reconstruction microscopy (STORM) have revealed TRF2-dependent T-loop formation²¹. Recently, nanopore sequencing has been developed to profile telomeric sequences^4,30,31. These structural insights have greatly enhanced our understanding of telomeric protein-DNA interactions. To further explore the dynamics of telomeric protein-DNA interactions, the development of new technologies is essential.

Single-molecule tools are powerful techniques for exploring protein-DNA interactions at telomere^32,33,34. Single-molecule mechanical methods, such as magnetic tweezers, optical tweezers and AFM, have been employed to investigate TRF2-dependent DNA distortion, reveal TRF2-mediated columnar stacking of human telomeric chromatin and observe processive telomerase catalysis, among other applications^{35,36,37,38,39,40}. These methods are particularly useful for probing topological conformations and the kinetics of protein-DNA association and dissociation.

However, the preparation of single-molecule constructs with telomeric repetitive motifs still presents challenges, which limits studies using single-molecule mechanical methods. To address this limitation, we have developed a single-molecule mechanical method to study global protein-DNA interactions on full-length human telomeres⁴¹. This method directly extracts telomeric DNA from human cells, circumventing the laborious preparation of artificial telomeric DNA. It facilitates the investigation of kinetic processes on long native telomeres spanning several kilobases.

In this protocol, we provide a detailed description of the steps for probing telomeric protein-DNA interactions using magnetic tweezers, a popular single-molecule mechanical tool^42,43,44. We demonstrate how to express and purify telomeric proteins, using TRF2 as an example, and how to prepare telomeric DNA from human cells. Additionally, we show how to set up a single-molecule assay on magnetic tweezers to study telomeric protein-DNA interactions, and we cover the subsequent data analysis of single-molecule experiments. This protocol will benefit researchers in the field of telomere biology and telomere-targeted drug discovery.

Protocol

1. General materials and methods

1. Refer to the Table of Materials file for the salts, chemicals, antibiotics, enzymes, antibodies, and resin materials used in this protocol.
2. Prepare Luria-Bertani (LB) liquid medium, LB agar plates, HEPES buffer, SDS polyacrylamide gel electrophoresis (SDS-PAGE), Phosphate-buffered saline (PBS), Tris-EDTA (TE) buffer according to the recipes from cold spring harbor protocols^{45,46,47,48,49,50,51,52,53}.
3. Acquire the pET28a vector (Addgene), prokaryotic, and eukaryotic cell lines (ATCC) through commercial sources. Then, continuously culture and passage them in the laboratory.
4. For large volumes of liquid, autoclave at 120 ˚C for 20 min. For small volumes, use a sterile filter with a 0.22 µm pore size.
5. Adjust the pH of the solutions to the desired level using HCl or NaOH before bringing them to their final volumes.
   NOTE: Magnetic tweezers are custom-built and run in an environment of LabVIEW 2017, while single-molecule data analysis is performed in MATLAB 2017^2,54,55.
6. For safety reasons, wear long-sleeved lab coats, nitrile gloves (or gloves made from a material that is impermeable and resistant to the substance), safety glasses or goggles, and closed-toe shoes.

2. Protein expression and purification of telomeric DNA-binding proteins

1. Perform cell culture and induce protein expression.
   1. Insert the coding sequence for the telomeric DNA-binding proteins (Figure 1A), TRF2 as the example in this case, into the pET28a vector by enzyme digestion and ligation, with SUMO employed as a fusion tag. Insert between the restriction sites of BamHI and HindIII, yielding the plasmid of pET28a-SUMO-TRF2 (Figure 1B).
   2. Transform BL21(DE3) cells with the pET28a-SUMO-TRF2 plasmid.
      1. Thaw a 50 µL aliquot of competent BL21(DE3) cells on ice.
      2. Add 1 µL of DNA of the pET28a-SUMO-TRF2 plasmid (containing 50 ng of DNA) to the competent cells. Gently swirl to mix without pipetting up and down.
         NOTE: Do not vortex.
      3. Incubate the mixture on ice for 30 min.
      4. For heat shock, heat the cell mixture at 42 °C for exactly 45 s in a water bath. Immediately return the cells to ice for 2 min to stabilize the transformed cells.
      5. Add 950 µL of room temperature (RT) LB medium to the cells to facilitate recovery.
      6. Incubate the transformed cells at 37 °C for 1 h while shaking at 220 rpm.
      7. Spread 100 µL of the transformation mixture on an LB agar plate containing 50 µg/mL kanamycin (85.8 µM).
      8. Incubate the plated cells overnight at 37 °C. Following incubation, select distinct colonies for use in the inoculation of starter cultures.
         NOTE: Colonies on plates can be stored at 4 °C for up to 2 weeks.
   3. Pick up a colony of the transformed BL21(DE3) cells and culture it in 5 mL of LB medium supplemented with 50 µg/mL kanamycin (85.8 µM). Incubate for 18 h at 37 °C with shaking at 220 rpm.
   4. To scale up, transfer 2 mL of the overnight culture to 200 mL of LB medium containing 50 µg/mL kanamycin (85.8 µM). Continue incubating at 37 °C with shaking at 220 rpm until the optical density at 600 nm (OD 600) reaches 0.6-0.8.
   5. Take 1 mL of the culture as an uninduced control sample.
   6. Induce the remaining culture with Isopropyl β-d-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM and incubate at 20 °C for 17 h to promote protein expression.
   7. Perform SDS-PAGE using a 5% stacking gel and a 10% separation gel. Load samples with SDS-PAGE loading buffer and run SDS-PAGE initially at 100 V for 30 min, followed by 120 V for 50 min in a Tris-Glycine buffer. Stain with Coomassie blue to visualize protein expression (see recipes in Supplementary Table 1) (Figure 1C).
      NOTE: See Discussion for troubleshooting if expression is not observed.
2. Purify the protein.
   1. Harvest the cells by centrifugation at 4 °C, 8500 x g for 10 min. Discard the supernatant and wash the pellet in 200 mL of PBS. Centrifuge again, discard the supernatant, and resuspend the cell pellet in 25 mL of lysis buffer (20 mM HEPES, 300 mM NaCl, 20 mM imidazole, 10% glycerol, 0.5 mM DTT, 1 mM phenylmethylsulfonyl fluoride [PMSF]).
      CAUTION: PMSF is corrosive and toxic and causes burns. Wear suitable protective clothing, gloves, and eye/face protection.
      ​NOTE: Freeze at -80 °C if not proceeding immediately.
   2. Add 5 µL of 100 mg/mL lysozyme, 5 µL of 1 U/µL DNase I, and 5 µL of 10 mg/mL RNase A to the resuspended cells.
   3. Perform sonication on ice, with 1 s bursts at 250 W followed by a 2 s pause, for a total of 30 min.
   4. Centrifuge at 38,000 x g, 4 °C for 40 min (30-60 min is acceptable) and filter the supernatant through a 0.22 µm syringe filter.
   5. Prepare 500 mL of Ni-column binding buffer by dissolving 8.76 g of NaCl (final concentration 300 mM) and 0.68 g of imidazole (final concentration 20 mM) in 20 mM HEPES (pH 8.0). Filter the solution through a 0.22 µm filter.
      CAUTION: Imidazole is dangerous and can cause skin and eye irritation. It may also harm an unborn child. Avoid contact with eyes, skin, and clothing. Do not ingest or inhale. Wear personal protective equipment, including face protection.
   6. Prepare Ni-column elution buffers (20 mM HEPES (pH 8.0), 300 mM NaCl) with increasing concentrations of imidazole (100 mM, 200 mM, 300 mM, and 500 mM) for stepwise elution, ensuring all buffers are appropriately filtered.
   7. Load the filtered supernatant onto the pre-equilibrated Ni-column in binding buffer (20 mM HEPES (pH 8.0),300 mM NaCl,20 mM imidazole). Wash with binding buffer and elute the protein with the prepared imidazole gradient, collecting 12 fractions, each with 1 mL (Figure 1C). Combine fractions with >95% purity.
   8. Measure the concentration of the eluted 6xHis-sumo-TRF2 fusion protein by A₂₈₀ in a spectrophotometer.
   9. For the protein used in single-molecule assays, add sumo protease according to the manufacturer's instructions (typically 0.125 U of protease per 2 µg of fusion protein) and allow overnight digestion at 4 °C.
   10. After sumo protease treatment, load the mixture back onto a Ni-column to bind any undigested fusion protein and the His-tagged sumo, allowing the tag-free TRF2 protein to flow through. Collect the flow-through.
3. Determine protein concentration and store the protein.
   1. Concentrate the purified TRF2 protein using a 30 kDa cutoff centrifugal filter unit and exchange it into a storage buffer containing 20 mM HEPES (pH 8.0), 150 mM NaCl, 0.5 mM DTT, to reduce the imidazole concentration below 150 mM.
   2. Repeat the concentration process until the volume is reduced to less than 1 mL.
   3. Analyze samples from each purification step by SDS-PAGE to assess the purity and integrity of the protein (Figure 1C).
      NOTE: Affinity chromatography is used to purify proteins, collect and wash eluted samples, and conduct SDS-PAGE to assess protein integrity and ensure quality control. When affinity chromatography alone does not meet purity requirements, size exclusion chromatography is employed to further enhance protein purity. UV absorbance curves help assess elution fractions, and SDS-PAGE verifies protein integrity, ensuring stringent quality control throughout the process.
   4. Add glycerol to a final concentration of 50% to the concentrated and buffer-exchanged protein for storage.
   5. Store the protein at -80 °C.

3. Preparation of human telomeric restriction fragments

1. Extract genomic DNA.
   1. Following the flowchart (Figure 2A), centrifuge 1 x 10⁷ cells at 1000 x g for 3 min and discard the supernatant. For washing, resuspend the cells in 200 µL of PBS, centrifuge again at 1000 x g for 3 min, and discard the supernatant.
   2. Add 760 µL of buffer containing 50 mM Tris-HCl (pH 8.0), 100 mM NaCl, 100 mM EDTA, and 1% SDS to the tube containing 1 x 10⁷ cells and gently resuspend by pipetting to avoid air bubbles.
      ​NOTE: Do not vortex.
   3. For the digestion of proteins and the elimination of DNases and RNases in the sample, add Proteinase K to a final concentration of 0.5 mg/mL, corresponding to 40 µL of a 10 mg/mL stock solution. Tap the bottom of the tube to mix (do not vortex).
   4. Incubate overnight at 37 °C.
   5. Add 265 µL of 5.4 M NaCl. Mix for 5 min by inverting the tube five times every minute, then place on ice for 20 min.
   6. Centrifuge at maximum speed (e.g., 16,900 x g) for 10 min at RT.
   7. Transfer the supernatant to a centrifuge tube and add an equal volume of isopropanol (approximately 750 µL), avoiding floating lipids or sediment. The DNA is in the supernatant.
   8. Invert the tube five times to mix. Visually confirm the presence of a pellet of DNA precipitating in the centrifuge tube.
   9. Centrifuge at RT at maximum speed for 10 min.
   10. Discard the supernatant. Wash the DNA pellet with 500 µL of 70% ethanol by inverting the centrifuge tube five times.
       NOTE: The pellet is DNA.
   11. Centrifuge at RT at maximum speed (e.g., 16,900 x g) for 10 min.
   12. Carefully remove all the supernatant, leaving the DNA pellet. Let the pellet air dry at RT for 5 min.
       NOTE: Avoid overdrying, as genomic DNA can become difficult to dissolve.
   13. Resuspend the DNA pellet in 475 µL of TE buffer (10 mM Tris-HCl [pH 8.0], 1 mM EDTA). Gently mix by tapping the bottom of the tube. For the digestion of RNA during DNA preparation, add 25 µL of 10 mg/mL RNase A (final concentration 0.5 mg/mL) and mix by gently tapping until the DNA pellet has completely dissolved.
       NOTE: Avoid vortexing to prevent DNA shearing.
   14. Incubate the DNA sample at 4 °C overnight.
   15. Perform an equal volume Tris-saturated phenol extraction: add 500 µL of phenol to the centrifuge tube, mix gently with a pipet, and centrifuge at maximum speed for 10 min at 4 °C.
       CAUTION: Exposure to phenol may cause irritation to the skin, eyes, nose, throat, and nervous system. Avoid contact with eyes, skin, and clothing. Do not ingest or inhale. Wear personal protective equipment, including face protection.
       NOTE: DNA is in the upper aqueous phase, proteins are in the interphase, and the organic phase is at the bottom.
   16. Repeat three times. Take the supernatant of DNA (approximately 280 µL).
   17. According to the volume of the DNA sample, add 1/10 volume (28 µL) of 3 M sodium acetate (final concentration 0.3 M, pH 5.2) and 2 volumes (616 µL) of cold 100% ethanol. Place the centrifuge tube at -80 °C for 2-3 h.
   18. Centrifuge at maximum speed (e.g., 16,900 x g) for 10 min at 4 °C. Thaw the solution on ice if frozen before centrifugation.
   19. Discard the supernatant and wash 3-4 times with 70% ethanol by inverting the centrifuge tube. Centrifuge at maximum speed for 5 min at 4 °C.
   20. Remove the ethanol and allow the pellet to air dry at RT for 5 min.
   21. Add 100 µL of TE buffer, tap the tube to mix, and let it sit at 4 °C for 2 h to fully dissolve.
       NOTE: Examine the integrity of the genomic DNA on a 1% agarose gel (Figure 2B).
   22. Store the DNA at -20 °C in the freezer. It will remain stable for at least 1 year.
2. Perform genomic DNA digestion and modification.
   1. Take 4 µg of genomic DNA and combine it with 1 µL of CviAII (10 U), 2 µL of 10x digestion buffer (see Supplementary Table 1 for recipes), and add water to reach a total volume of 20 µL. Incubate the mixture at 25 °C for 12 h.
      NOTE: FatI can replace CviAII, which is now discontinued by NEB.
   2. Add 1 µL of NdeI (20 U), 1 µL of MseI (10 U), and 1 µL of BfaI (10 U) to the same tube containing the previous mixture (20 µL). Also, add 2 µL of 10x digestion buffer and 15 µL of water to achieve a total volume of 40 µL. Incubate at 37 °C for 12 h, then heat-inactivate all enzymes at 80 °C for 20 min.
      ​NOTE: The digestion of genomic DNA can be examined using the Terminal Restriction Fragment (TRF) method (Figure 2C). The combination of the four restriction enzymes (CviAII, NdeI, MseI, and BfaI) digest the genomic DNA into fragments of about 800 bp and yields the TRFs with minimal subtelomeric DNA contamination.
   3. For Digoxigenin modification, take 40 µL of the digested genomic DNA and add 4 µL of 1 mM dATP (final concentration 0.08 mM), 4 µL of 1 mM digoxigenin-11-dUTP (final concentration 0.08 mM), 1 µL of Klenow Fragment (5 U), 0.5 µL of 100 mM DTT (final concentration 1 mM), and 1 µL of 10x digestion buffer. Incubate at 37 °C for 12 h, followed by heating at 75 °C for 20 min to inactivate the Klenow Fragment.
   4. For biotin labeling, take 12.5 µL of the Digoxigenin-modified genomic DNA (1 µg) and add 1 µL of 1 µM biotinylated telomeric probe (i.e., 1 pmol) that are complementary to the telomeric single-stranded overhang. Add 611.5 µL of TE Li buffer (10 mM Tris-HCl (pH 8.0), 1 mM EDTA, 50 mM LiCl) to dilute the 50 mM K ions to 1 mM, making a total volume of 625 µL, which can be divided into 16 tubes with 39 µL each.
   5. Heat at 75 °C for 3 min in a thermal cycler, then decrease the temperature by 0.1 °C every 30 s until reaching 25 °C.
   6. Store the DNA samples at -20 °C for at least a year.

4. Setting up a flow cell for telomeric DNA sample on magnetic tweezers

1. Make a flow cell for single-molecule assays.
   1. Use an electric grinder to drill holes as the inlet and outlet on the top coverslips (# 2 cover glass with a thickness of 0.2 mm).
   2. Clean the coverslips in detergent by sonication for 30 min.
   3. Wash the cover slips with ultra-pure water 3 times.
   4. Clean the coverslips in isopropanol by sonication for 30 min.
   5. Wash the coverslips with ultra-pure water 3 times.
   6. Clean the coverslips in ultra-pure by sonication for 15-30 min.
   7. Dry the coverslips with nitrogen.
      NOTE: The protocol can be stopped here and the cleaned coverslips can be stored for later use.
   8. Coat the bottom coverslips (without holes) with 20 µL of 0.1% nitrocellulose and add reference beads (20x dilution, 3 µm diameter). Bake at 100 °C for 4 min.
   9. Use a scalpel to cut double-layer parafilm spacers according to a metal mold. The mold is a rectangular aluminum alloy piece with the same dimensions as the coverslip, featuring a hollowed-out center to facilitate channel cutting.
   10. Assemble the flow cell sandwich (Figure 3A). Heat at 85 °C and use two swabs to massage until the parafilm seals the channel.
   11. Coat the flow cell with an antibody. Inject 70 µL of anti-digoxigenin antibody (0.1 mg/mL) directly into the flow cell. Leave at RT for 1 h.
   12. Passivize the flow cell. Inject 70 µL of 10 mg/mL BSA to displace the anti-digoxigenin antibody. Leave at RT for 12 h. Unbound anti-digoxigenin antibodies will be flushed away in the following steps with rigorous flushing.
       NOTE: Store the flow cell at 4 °C for 1-2 weeks.
   13. Test for non-specific binding by flushing 5 µL of washed MyOne (10 mg/mL) (or 5 µL of washed M270 (10 mg/mL)) beads without DNA into the flow cell. Leave for 10 min, then flush beads away using a working buffer (20 mM HEPES (pH 7.5), 100 mM NaCl, 1 mM EDTA). Count the number of beads stuck to the surface in one field of view. A well-treated surface should show zero or only a few beads.
2. Isolate and immobilize telomeric DNA tethers in a flow cell.
   1. Take 10 µL of M270 beads (10 mg/mL) or 5 µL of 10 mg/mL MyOne and wash them five times with 50 µL of working buffer (20 mM HEPES [pH 7.5], 100 mM NaCl, 1 mM EDTA) using a magnet.
   2. Take 500 ng of digested genomic DNA and put it into a 1.5 mL centrifuge tube. Add all the beads (50 µL) on top of the DNA sample.
   3. Without pipetting, gently flick the centrifuge tube a few times to mix the beads and DNA sample. Leave the mixture on ice for 1 h. Wash with 500 µL of working buffer three times using a magnet to pull the beads down, with 10 min intervals between washes.
   4. Resuspend the sample using 30 µL of working buffer and load the mixture into the flow cell. Leave for 30 min. Flush unbound magnetic beads.
      NOTE: Telomeric DNA is tethered between the bottom cover slip and the magnetic bead through affinity interactions mediated by digoxigenin-antibody and biotin-streptavidin (Figure 3B).
3. Set up a flow cell on magnetic tweezers
   1. Clean the flow cell with 70% ethanol and dry the surface using lens tissue. Place the cleaned flow cell on the lower sample holder and gently assemble the upper sample rack using a screwdriver.
   2. Apply a drop of lens oil to the bottom surface of the flow cell, where it aligns with the objective lens. Then, place the sample holder on top of the objective lens and secure it using a screwdriver.
   3. Select a pair of 5 mm cubic magnets arranged in a vertical configuration. Align the magnet holder with the X-axis of the magnetic tweezers' light path for imaging.
      ​NOTE: This is a critical step. The direction of the magnetic field is aligned and thus determined by the orientation of the magnet holder.
   4. Launch the graphical programming software and connect the controllers for the magnetic tweezers. Adjust the field of view to locate a reference bead at the bottom of the flow cell and adjust the objective lens slightly so that the reference bead shows clear diffraction rings.
   5. Move the magnets down to the top of the flow cell.
      NOTE: This is a critical step. Sudden changes in the bead diffraction pattern indicate that the magnets have just touched the surface of the flow cell. This position is considered the offset for the measurement zero point.
      NOTE: Lower the magnets carefully, starting with larger steps and gradually shifting to smaller steps. Move in 0.1 mm increments to avoid damaging the flow cell when approaching the flow cell.
   6. Raise the magnets to their highest position and remove the magnet holder.
   7. Turn on the peristaltic pump connected to the waste bottle and discard the liquid. Add 100 µL of working buffer to the inlet and set the peristaltic pump flow rate to 100 µL/min to fill the flow cell with buffer.

5. Measurements of a telomere using single-molecule magnetic tweezers

1. Set up the camera.
   1. Set the grayscale of the complementary metal-oxide semiconductor (CMOS) camera with a brightness adjustment to 150 levels.
   2. Define the size of the region of interest (ROI).
   3. Set the framerate to 200 Hz with an exposure time of 5000 µs.
      NOTE: This is a critical step. Ensure the shutter dead time is set to zero.
   4. Move the magnet position to 3 cm (approximately 8 pN) and adjust the objective focus on the beads.
2. Establish the look-up table (LUT).
   1. Move the magnets to a position corresponding to 8 pN to tightly stretch the DNA-tethered magnetic bead.
   2. Set the LUT to 200 steps with a step size of 0.05 µm.
   3. Use a CMOS camera to capture the profiles of the magnetic beads at different positions to establish the LUT.
      ​NOTE: This is a critical step. Polystyrene beads fixed in the flow cell serve as reference beads to eliminate drift for the beads of interest.
3. Design a single-molecule mechanical experiment.
   1. Write a script in MATLAB to control motor movements for force ramp or force jump assays.
      NOTE: This is a critical step. This script encodes the commands for magnet movements. The force loading rate is set at this step for a force ramp assay. The levels and durations of force jump assays are also determined here (Figure 4, Supplementary File 1, Supplementary File 2, and Supplementary File 3).
   2. Import the script into graphical programming software to test the single-molecule experiments.
   3. Check the total frames required for running the single-molecule experiment and ensure that the available memory exceeds this requirement.
      NOTE: Data will be lost if the allocated memory is less than required because saving data takes time.
   4. Verify the maximum imaging speed to determine the maximum number of beads that can be monitored simultaneously.
      NOTE: The experiment will be terminated if the imaging speed limit is exceeded.
   5. Using a force jump assay as an example, run the single-molecule mechanical experiment. Forces are abruptly applied to the magnetic bead, which transmits these forces to the DNA molecule, immediately stretching it upwards.

6. Measurements of TRF1/2 on a telomere using magnetic tweezers

1. Force ramp assay
   1. Using TRF1 as an example, load 200 µL of 10 nM TRF1 into the flow cell at a slow flow rate (e.g., 30 µL/min). Allow 30 min for TRF1 to bind to the telomeric DNA.
   2. Choose a script for a force ramp experiment with a force loading rate of ±1 pN/s.
   3. Name the data files appropriately and run the experiment. The data will be saved to the specified destination for analysis (Figure 5).
      NOTE: During this experiment, the force is linearly increased and decreased between 0 pN and 17 pN, which stretches and relaxes the telomeric DNA and breaks the DNA loops mediated by TRF1/2.
2. Force Jump assay
   1. Select a script to run a force jump assay.
      NOTE: Various forces are applied to stretch the telomeric DNA, for example, "Rest force" (Frest) at 0 pN for 40 s, "Test force" (Ftest) ranging from 2-8 pN for 60 s, "High force" (Fhigh) at 10 pN for 30 s, and "Maximum force" (Fmax) at 20 pN for 30 s.
   2. Name the data files appropriately and run the experiment. The data will be saved to the specified destination for analysis (Figure 5).

Results

Figure 1A illustrates the schematic domains and structures of TRF1 and TRF2, consisting of 439 and 542 amino acids, respectively, which can be expressed in prokaryotic cells. The preparation of TRF1 has been previously described in the literature⁴¹. Here, we provide a comprehensive description and representative results of the preparation of TRF2. Figure 1B shows the plasmid map used for expressing TRF2 in E. coli. We evaluated TRF2 expression before and after induction in E. coli, as depicted in Figure 1B. The purification and tag cleavage processes were also analyzed using SDS-PAGE (Figure 1C). Following this protocol, both telomeric binding proteins, TRF1 and TRF2, were successfully expressed in E. coli.

Telomeric DNA, in the form of terminal restriction fragments (TRFs), is generated from human genomes using a combination of four restriction enzymes. Following the flowchart shown in Figure 2A, we extracted genomes from human cells. The enzymes selectively digest genomic DNA while leaving the full-length telomeric DNA intact, thus providing material for single-molecule assays. We have extensively tested the TRF DNA preparation using various human cell lines. The integrity of the genomic DNA was examined using agarose gel electrophoresis, as shown in Figure 2B. The TRFs were subsequently analyzed by Southern blotting, employing fluorescently labeled oligonucleotides complementary to the telomeric repeat sequence TTAGGG (Figure 2C), following a method described elsewhere^41,56. On average, the length of human TRFs is around a few kilobases.

Single-molecule assays take place in a flow cell assembled as described in Figure 3A. A nitrocellulose coating covers the negatively charged glass surface, providing a hydrophilic matrix for antibody adsorption. BSA passivation blocks areas of the nitrocellulose matrix not bound by anti-digoxigenin antibodies. Polystyrene beads, which do not contain iron nanoparticles, serve as reference beads. TRF tethers are formed through affinity interactions between the digoxigenin-antibody on the nitrocellulose matrix and biotin-streptavidin on the magnetic bead surface (Figure 3B). TRF1 or TRF2 are introduced into the flow cell, binding to the TRF DNA tethers via free diffusion. Magnetic tweezers are used to modulate the magnetic field, generating forces on the magnetic beads to stretch and relax the TRF tethers, thereby enabling the probing of protein-DNA interactions.

A single-molecule mechanical assay is designed by developing a script to control the motions of the magnetic tweezers' motors. A flowchart provides a structured approach to creating this script in MatLab for a single-molecule assay using magnetic tweezers (Figure 4A). The magnets move along the z-axis within a range of 0-25 mm. Forces are calculated based on the magnet positions, according to the magnet configuration, using equations described in the literature⁵⁴. For a specific force-loading rate, the magnet positions and movement speeds are designed and programmed in MatLab to achieve the desired force manipulation profile, as demonstrated in the force ramp assay example (Figure 4B-D).

Using TRF1 as an example, we probed the telomeric DNA-protein interactions with single-molecule magnetic tweezers. The experimental setup can be configured as a force-ramp assay, as shown in Figure 5A, where the force-extension curves exhibit zigzag features during stretching due to the breaking of protein-DNA interactions and smooth traces during relaxation, reflecting the DNA fiber without protein-mediated loops. The setup can also be configured as a force-jump assay, as shown in Figure 5B,C, allowing us to measure changes in extension and the durations of protein-DNA interactions under specific forces. The dissociation kinetics of the telomeric DNA-protein complexes can be derived from these measurements (Figure 5D). Additionally, the formation of loops in the DNA-protein complexes can be revealed by changes in extension (Figure 5E). Furthermore, the protein concentration can be titrated within this telomeric DNA experimental setup. Moreover, the length heterogeneity of telomeric DNA from human cells allows us to investigate the loop formation mechanism in telomeres of various lengths (Figure 6).

Figure 1: The expression and purification of TRF1/2. (A) The domain architecture of TRF1 and TRF2. (B) The plasmid map of pET28a-SUMO-TRF2 for expressing TRF2 in E. coli. (C) SDS-PAGE analysis of the samples obtained at various stages of expression and purification. Lane 1: Uninduced by IPTG. Lane 2: IPTG-induced precipitate obtained by cell lysis. Lane 3: IPTG-induced supernatant obtained after cell lysis. Lane 4: Flow through Ni column. Lane 5: Wash with low imidazole buffer to remove impurity proteins. Lane 6: Elution of 6xHis-SUMO-TRF2 with buffer containing 300 mM imidazole. Lane 7: Digestion of SUMO tags by SUMO protease enzyme. Lane 8: Repurification with Ni column and elution of TRF2 (6xHis and SUMO tags removed) with buffer containing 20 mM imidazole. Please click here to view a larger version of this figure.

Figure 2: Preparation of terminal restriction fragments (TRF). (A) Flowchart for the preparation of TRFs from human cells. (B) Examination of genomic DNA integrity using a 1% agarose gel for 5 human cell lines. (C) TRF analysis performed by Southern blotting for 5 human cell lines. Please click here to view a larger version of this figure.

Figure 3: Preparation of single-molecule mechanical assays. (A) Flowchart depicting the preparation of a flow cell for single-molecule mechanical assays. (B) Schematic representation of single-molecule mechanical assays using magnetic tweezers. Please click here to view a larger version of this figure.

Figure 4: Design of a single-molecule mechanical assay. (A) Flowchart for generating a script to execute a force-ramp assay. (B) The movement profile of magnets along the z-axis. (C) The force profile corresponding to magnet movement. (D) The correlation between the magnet positions and the resulting forces. Please click here to view a larger version of this figure.

Figure 5: Telomeric DNA-protein interactions probed by single-molecule magnetic tweezers. (A) TRF1-telomere interactions were probed in force-ramp assays (N = 10) in a buffer containing 20 mM HEPES (pH 7.5), 1 mM EDTA, and 100 mM NaCl at 23 °C. The concentration of TRF1 was 10 nM, with a force loading rate of ±1 pN/s. (B) TRF tether was stretched and relaxed in force-jump assays. The applied force protocol was Frest = 0 pN, Ftest = 2 pN - 8 pN, Fhigh = 10 pN, and Fmax = 20 pN. Buffer and temperature conditions were the same as in (A). Frame rate = 200 Hz. (C) TRF1-telomere interactions were probed in force-jump assays with a TRF1 concentration of 10 nM. (D) Dissociation kinetics of TRF1-telomere complexes. The logarithm of the dissociation rate at tested forces (Mean ± SD, n = 206) follows the Kramer-Bell-Evans model (red curve and legend equation). The inset shows the average dissociation time (<τ>) at Ftest. (E) Distribution of changes in extension (ΔL) upon rupture events of TRF1-telomere complexes. Black and red curves represent Gaussian fittings. This figure has been modified with permission from Li et al.⁴¹. Please click here to view a larger version of this figure.

Figure 6: The number and sizes of DNA loops formed by TRF1 in a telomere. (A) Telomere length dependency of DNA loop sizes (ΔL) at [TRF1] = 20 nM. Curves represent Gaussian fittings. (B) Telomere length dependency of ΔL at [TRF1] = 40 nM. (C) Correlation between ΔL and the number of rupture events per telomere, N. The zero-order correlation is r = -0.20, with p < 0.001 (sample size = 1496). (D) A cartoon illustrating a possible mechanism explaining the negative correlation between ΔL and N suggesting that TRF1 can compact a single telomere with primary loop domains into a high-order topology. This figure has been modified with permission from Li et al.⁴¹. Please click here to view a larger version of this figure.

Supplementary File 1: MatLab code for generating scripts.m Please click here to download this File.

Supplementary File 2: Script for constant force assay.txt. Please click here to download this File.

Supplementary File 3: Script for force ramp assay.txt. Please click here to download this File.

Supplementary Table 1: Recipes Please click here to download this File.

Discussion

This protocol employs magnetic tweezers for the manipulation of TRFs at the single-molecule level^57,58,59. We utilize magnetic beads to separate TRFs from genomic DNA fragments. Following restriction digestion, TRFs bind to the magnetic beads, enabling their easy separation from genomic DNA fragments. This approach allows for manipulation using magnetic tweezers, which can effectively trap magnetic beads, unlike optical tweezers that are limited by transparency issues. Moreover, while AFM can be used for single-molecule manipulation, additional steps are required to dissociate TRFs from magnetic beads and immobilize them between a mica surface and a cantilever tip. In contrast, magnetic tweezers offer a more straightforward and efficient method for single-molecule manipulation of TRFs, eliminating the need for these extra steps⁴¹.

Maintaining the integrity of genomic DNA and TRFs is crucial, as single-molecule manipulation depends on intact TRFs. Using magnetic tweezers, we stretch and relax TRFs by holding their two ends to probe protein-DNA interactions. Accurate force measurement on magnetic beads is fundamental for single-molecule mechanical assays⁵⁴. Thus, cubic-shaped magnets should be aligned parallel or orthogonal to the camera's light path in magnetic tweezers (NOTE below step 4.3.3), ensuring precise three-dimensional bead position recording per established equations^54,60. Magnet offset is crucial for determining their positions and the resulting forces on magnetic beads. We use #2 cover glass with a thickness of approximately 0.2 mm for consistency or adjust the offset in the equation for different cover glasses⁵⁴. Frame rate is vital for capturing the dynamics of protein-DNA interactions, necessitating a shutter dead time of zero for complete bead movement data. The motor movements, encoded by a script, are central to designing single-molecule mechanical assays. Critical parameters such as force loading rate, forces, and durations should be set and optimized iteratively. Protein concentrations used in single-molecule assays require extensive titration to determine the optimal level for studying protein-DNA interactions. Initial force testing can be performed using force-ramp assays with force spectroscopy, while force-jump assays evaluate forces of interest at a constant level.

One may encounter a few challenges when adapting this protocol. If TRF1 or TRF2 is not expressed in E. coli, start with a small-scale trial, lyse cells, and analyze proteins via SDS-PAGE and staining, confirming with Western Blot using tag-specific antibodies. One should verify the construct and plasmid to ensure correct cloning without mutations, frameshifts, or premature stop codons, and check promoter suitability for E. coli. The expression should be optimized by testing different E. coli strains (e.g., BL21(DE3), Rosetta), adjusting growth temperature (15-25 °C), induction time (2-16 h), and IPTG concentration (0.1-1 mM). Inclusion bodies should be addressed by analyzing cell pellets before and after sonication. One should tackle codon bias with strains providing rare tRNAs or codon optimization. Try different growth media (LB, TB, auto-induction) and improve aeration by adjusting shaking speed and culture volume. For toxicity issues, use tightly regulated promoters or lower-copy-number vectors and add glucose (0.2%) to repress leaky lac promoter expression. Maintain plasmid integrity with proper antibiotic selection and ensure complete protein release by optimizing the cell lysis method^61,62.

If no TRF tethers are found in single-molecule assays, one possibility is that not enough initial genomic DNA was used. Try using 500 ng of genomic DNA equivalent of TRFs to set up a single-molecule mechanical assay. Persistent issues with finding TRF tethers in magnetic tweezers might be due to excessively long digestion times of genomic DNA. Keep the digestion procedure under 12 h to avoid star activity. Additionally, the formation of G-quadruplexes at the single-stranded overhang of telomeric DNA could inhibit the annealing of biotin-labeled oligonucleotides for binding to magnetic beads. To prevent this, maintain potassium or sodium ion concentrations below 1 mM to avoid G-quadruplex formation and increase the likelihood of forming TRF tethers in the single-molecule setup. Additionally, increasing the heating temperature during the annealing step (step 3.2.5) helps to destabilize G-quadruplexes and facilitates the binding of biotin-labeled oligonucleotides to TRFs.

In the presence of TRF1 or TRF2 proteins, TRF DNA tethers could become tightly compacted and too difficult to stretch in magnetic tweezers due to the abundance of binding sites on the telomere for these proteins. Titrating the protein concentrations can help identify optimal levels for single-molecule mechanical assays. The protein-DNA binding affinity can be assessed using EMSA or SPR assays to determine the dissociation constant (K_d), which can then inform suitable concentrations for single-molecule assays.

The limited amount of TRFs generated from genomic DNA could hold back the efficiency of single-molecule assays. Consequently, collecting single-molecule data may take longer time compared to using artificially synthesized telomeric DNA constructs. However, using telomeres sourced directly from the human genome provides several advantages that artificial telomeric DNA, commonly prepared by plasmid in bacteria or PCR methods, cannot offer. One advantage is that human TRFs are heterogeneous in length, allowing the examination of telomere length dependency in protein-DNA interactions. In contrast, artificial telomeric DNA is usually uniform in length, and generating artificial telomeric DNA of various lengths would be time-consuming and labor-intensive. Additionally, human TRFs contain original sequences from cells, including natural markers such as modifications, mutations, and damages. This allows for the investigation of protein-DNA interactions under physiological or pathological conditions, which is not possible with artificial telomeric DNA.

This protocol is essential for studies in the field of telomere biology at the single-molecule level and can potentially benefit telomere-targeting drug discovery.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-Digoxigenin	Roche	11214667001
BfaI	New England Biolab (NEB)	R0568S
BSA	Sigma-Aldrich	V900933
CMOS camera	Mikrotron	MC1362
CviAII	New England Biolab (NEB)	R0640S
DIG-11-dUTP	Jena Bioscience	NU-803-DIGXL
DNA extraction solution	G-CLONE	EX0108
Dnase I, Rnase-Free, Hc Ea	Thermo Fisher Scientific	EN0523
dNTP mixture	Nanjing Vazyme Biotech Co., Ltd (Vazyme)	P032-02
DTT	Solarbio	D1070
Dynabeads M-270  beads	Thermo Fisher Scientific	65305	Streptavidin beads
Dynabeads MyOne beads	Thermo Fisher Scientific	65001	Streptavidin beads
Ethanol	Tianjin No.6 Chemical Reagent Factory	1083
Glycerol	Beijing Hwrkchemical Co,. Ltd	SMG66258-1
Imidazole	Solarbio	II0070
IPTG	Solarbio	I8070
Isopropanol	Tianjin No.6 Chemical Reagent Factory	A1079
Kanamycin	Thermo Fisher Scientific	EN0523
Klenow fragment (3′-5′ exo-)	New England Biolab (NEB)	M0212S
LabView	National Instruments	https://www.ni.com/en-us/shop/product/labview.html	Graphical programming software
LiCl	Bide Pharmatech Co., Ltd (bidepharm)	BD136449
Lysozyme	Solarbio	L8120-5
MseI	New England Biolab (NEB)	R0525S
NaCl	Shanghai Aladdin	C111533
NanoDrop	Thermo Fisher Scientific		Spectrophotometer
NdeI	New England Biolab (NEB)	R0111S
Ni NTA Beads 6FF	Changzhou Smart-Lifesciences Biotechnology Co.,Ltd	SA005025
Nitrocellulose membrane	ABclonal	RM02801
PMSF	Solarbio	P8340
Proteinase K	Beyotime Biotech Inc (beyotime)	ST535-500mg
rCutSmart Buffer	New England Biolab (NEB)	B6004S
Rnase A	Sigma-Aldrich	R4875
Sodium acetate	SERVA Electrophoresis GmbH	2124902
Sumo protease	Beyotime Biotech Inc (beyotime)	P2312M