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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

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 chromosomes1,2,3. Telomere erosion during cell division leads to cell senescence and aging, while abnormal elongation of telomeres contributes to cancer4,5. For telomeres to function properly, their replication, elongation, and damage responses must be highly regulated6,7,8. Shelterin, composed of six subunits, plays a central role in telomere protection9,10,11. A deeper understanding of telomeres will provide valuable insights into telomere biology.

TRF1 and TRF2, core subunits of shelterin, are telomeric binding proteins12,13. 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 proteins15,16,17,18,19. TRF2 is particularly important for the formation of telomeric D-loops and T-loops20,21. Due to their crucial roles in telomere protection, TRF1 and TRF2 have emerged as potential anti-cancer drug targets22,23,24,25.

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 affinities20,26. 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 sequences4,30,31. 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 telomere32,33,34. 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 applications35,36,37,38,39,40. 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 tool42,43,44. 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 protocols45,46,47,48,49,50,51,52,53.
  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 ¡m 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 20172,54,55.
  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 Β΅L aliquot of competent BL21(DE3) cells on ice.
      2. Add 1 Β΅L 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 Β°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 Β΅L of room temperature (RT) LB medium to the cells to facilitate recovery.
      6. Incubate the transformed cells at 37 Β°C for 1 h while shaking at 220 rpm.
      7. Spread 100 Β΅L of the transformation mixture on an LB agar plate containing 50 Β΅g/mL kanamycin (85.8 Β΅M).
      8. Incubate the plated cells overnight at 37 Β°C. Following incubation, select distinct colonies for use in the inoculation of starter cultures.
        NOTE: Colonies on plates can be stored at 4 Β°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 Β΅g/mL kanamycin (85.8 Β΅M). Incubate for 18 h at 37 Β°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 Β΅g/mL kanamycin (85.8 Β΅M). Continue incubating at 37 Β°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 Ξ²-d-1-thiogalactopyranoside (IPTG) to a final concentration of 1 mM and incubate at 20 Β°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 Β°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 Β°C if not proceeding immediately.
    2. Add 5 Β΅L of 100 mg/mL lysozyme, 5 Β΅L of 1 U/Β΅L DNase I, and 5 Β΅L 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 Β°C for 40 min (30-60 min is acceptable) and filter the supernatant through a 0.22 Β΅m 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 Β΅m 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 Β΅g of fusion protein) and allow overnight digestion at 4 Β°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 Β°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 Β΅L of PBS, centrifuge again at 1000 x g for 3 min, and discard the supernatant.
    2. Add 760 Β΅L 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 Β΅L of a 10 mg/mL stock solution. Tap the bottom of the tube to mix (do not vortex).
    4. Incubate overnight at 37 Β°C.
    5. Add 265 Β΅L 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 Β΅L), 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 Β΅L 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 Β΅L 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 Β΅L 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 Β°C overnight.
    15. Perform an equal volume Tris-saturated phenol extraction: add 500 Β΅L of phenol to the centrifuge tube, mix gently with a pipet, and centrifuge at maximum speed for 10 min at 4 Β°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 Β΅L).
    17. According to the volume of the DNA sample, add 1/10 volume (28 Β΅L) of 3 M sodium acetate (final concentration 0.3 M, pH 5.2) and 2 volumes (616 Β΅L) of cold 100% ethanol. Place the centrifuge tube at -80 Β°C for 2-3 h.
    18. Centrifuge at maximum speed (e.g., 16,900 x g) for 10 min at 4 Β°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 Β°C.
    20. Remove the ethanol and allow the pellet to air dry at RT for 5 min.
    21. Add 100 Β΅L of TE buffer, tap the tube to mix, and let it sit at 4 Β°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 Β°C in the freezer. It will remain stable for at least 1 year.
  2. Perform genomic DNA digestion and modification.
    1. Take 4 Β΅g of genomic DNA and combine it with 1 Β΅L of CviAII (10 U), 2 Β΅L of 10x digestion buffer (see Supplementary Table 1 for recipes), and add water to reach a total volume of 20 Β΅L. Incubate the mixture at 25 Β°C for 12 h.
      NOTE: FatI can replace CviAII, which is now discontinued by NEB.
    2. Add 1 Β΅L of NdeI (20 U), 1 Β΅L of MseI (10 U), and 1 Β΅L of BfaI (10 U) to the same tube containing the previous mixture (20 Β΅L). Also, add 2 Β΅L of 10x digestion buffer and 15 Β΅L of water to achieve a total volume of 40 Β΅L. Incubate at 37 Β°C for 12 h, then heat-inactivate all enzymes at 80 Β°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 Β΅L of the digested genomic DNA and add 4 Β΅L of 1 mM dATP (final concentration 0.08 mM), 4 Β΅L of 1 mM digoxigenin-11-dUTP (final concentration 0.08 mM), 1 Β΅L of Klenow Fragment (5 U), 0.5 Β΅L of 100 mM DTT (final concentration 1 mM), and 1 Β΅L of 10x digestion buffer. Incubate at 37 Β°C for 12 h, followed by heating at 75 Β°C for 20 min to inactivate the Klenow Fragment.
    4. For biotin labeling, take 12.5 Β΅L of the Digoxigenin-modified genomic DNA (1 Β΅g) and add 1 Β΅L of 1 Β΅M biotinylated telomeric probe (i.e., 1 pmol) that are complementary to the telomeric single-stranded overhang. Add 611.5 Β΅L 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 Β΅L, which can be divided into 16 tubes with 39 Β΅L each.
    5. Heat at 75 Β°C for 3 min in a thermal cycler, then decrease the temperature by 0.1 Β°C every 30 s until reaching 25 Β°C.
    6. Store the DNA samples at -20 Β°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 Β΅L of 0.1% nitrocellulose and add reference beads (20x dilution, 3 Β΅m diameter). Bake at 100 Β°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 Β°C and use two swabs to massage until the parafilm seals the channel.
    11. Coat the flow cell with an antibody. Inject 70 Β΅L 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 Β΅L 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 Β°C for 1-2 weeks.
    13. Test for non-specific binding by flushing 5 Β΅L of washed MyOne (10 mg/mL) (or 5 Β΅L 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 Β΅L of M270 beads (10 mg/mL) or 5 Β΅L of 10 mg/mL MyOne and wash them five times with 50 Β΅L 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 Β΅L) 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 Β΅L 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 Β΅L 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 Β΅L of working buffer to the inlet and set the peristaltic pump flow rate to 100 Β΅L/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 Β΅s.
      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 Β΅m.
    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 Β΅L of 10 nM TRF1 into the flow cell at a slow flow rate (e.g., 30 Β΅L/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 Β±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 T...

Discussion

This protocol employs magnetic tweezers for the manipulation of TRFs at the single-molecule level57,58,59. 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...

Disclosures

The authors declare no competing financial interests or other conflicts of interest.

Acknowledgements

This work was supported by the National Natural Science Foundation of China [Grant 32071227 to Z.Y.], Tianjin Municipal Natural Science Foundation of China (22JCYBJC01070 to Z.Y.), and State Key Laboratory of Precision Measuring Technology and Instruments (Tianjin University) [Grant pilab2210 to Z.Y.].

Materials

NameCompanyCatalog NumberComments
Anti-DigoxigeninRoche11214667001
BfaINew England Biolab (NEB)R0568S
BSASigma-AldrichV900933
CMOS cameraΒ MikrotronMC1362
CviAIINew England Biolab (NEB)R0640S
DIG-11-dUTPJena BioscienceNU-803-DIGXL
DNA extraction solutionG-CLONEEX0108
Dnase I, Rnase-Free, Hc EaThermo Fisher ScientificEN0523
dNTP mixtureNanjing Vazyme Biotech Co., Ltd (Vazyme)P032-02
DTTSolarbioD1070
Dynabeads M-270Β  beadsThermo Fisher Scientific65305Streptavidin beads
Dynabeads MyOne beadsThermo Fisher Scientific65001Streptavidin beads
EthanolTianjin No.6 Chemical Reagent Factory1083
GlycerolBeijing Hwrkchemical Co,. LtdSMG66258-1
ImidazoleSolarbioII0070
IPTGSolarbioI8070
IsopropanolTianjin No.6 Chemical Reagent FactoryA1079
KanamycinThermo Fisher ScientificEN0523
Klenow fragment (3β€²-5β€² exo-)New England Biolab (NEB)M0212S
LabViewNational Instrumentshttps://www.ni.com/en-us/shop/product/labview.htmlGraphical programming softwareΒ 
LiClBide Pharmatech Co., Ltd (bidepharm)BD136449
LysozymeSolarbioL8120-5
MseINew England Biolab (NEB)R0525S
NaClShanghai AladdinC111533
NanoDropThermo Fisher ScientificSpectrophotometer
NdeINew England Biolab (NEB)R0111S
Ni NTA Beads 6FFChangzhou Smart-Lifesciences Biotechnology Co.,LtdSA005025
Nitrocellulose membraneABclonalRM02801
PMSFSolarbioP8340
Proteinase KBeyotime Biotech Inc (beyotime)ST535-500mg
rCutSmart BufferNew England Biolab (NEB)B6004S
Rnase ASigma-AldrichR4875
Sodium acetateSERVA Electrophoresis GmbH2124902
Sumo proteaseBeyotime Biotech Inc (beyotime)P2312M

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