DNA Molecular Handles for Single-Molecule Protein-Folding Studies by Optical Tweezers

  • Ciro Cecconi
  • Elizabeth A. Shank
  • Susan Marqusee
  • Carlos Bustamante
Protocol
Part of the Methods in Molecular Biology book series (MIMB, volume 749)

Abstract

In this chapter, we describe a method that extends the use of optical tweezers to the study of the folding mechanism of single protein molecules. This method entails the use of DNA molecules as molecular handles to manipulate individual proteins between two polystyrene beads. The DNA molecules function as spacers between the protein and the beads, and keep the interactions between the tethering surfaces to a minimum. The handles can have different lengths, be attached to any pair of exposed cysteine residues, and be used to manipulate both monomeric and polymeric proteins. By changing the position of the cysteine residues on the protein surface, it is possible to apply the force to different portions of the protein and along different molecular axes. Circular dichroism and enzymatic activity studies have revealed that for many proteins, the handles do not significantly affect the folding behavior and the structure of the tethered protein. This method makes it possible to study protein folding in the physiologically relevant low-force regime of optical tweezers and enables us to monitor processes – such as refolding events and fluctuations between different molecular conformations – that could not be detected in previous force spectroscopy experiments.

Key words

Laser tweezers DNA handles Protein–DNA chimeras Single-molecule mechanical manipulation Protein folding 

1 Introduction

During the last decade, single-molecule force spectroscopy has emerged as a new and powerful technique to study protein folding. By manipulating one molecule at a time with either an atomic force microscope (AFM) or optical tweezers, scientists have been able to investigate aspects the folding mechanism of proteins that were previously inaccessible to experimental investigation, such as distances from the folded and unfolded states to their corresponding transitions states or anisotropy of a protein’s energy landscape (1, 2, 3, 4). The vast majority of these studies have been carried out by stretching engineered polymeric proteins between a gold substrate and an AFM silicon nitride tip (5). Using this method, scientists have applied AFM to the study of the unfolding processes of a large variety of proteins (6, 7). Optical tweezers instead have had much more limited use. The microscopic beads used in optical tweezers experiments are in fact not suited to manipulate molecules whose structures span only a few nanometers, such as most globular proteins and many of the polymeric proteins used in AFM studies; at such short distances, the surfaces of the tethering beads interact and severely compromise the measurements. For these reasons, optical tweezers have long been restricted to the characterization of the mechanical properties of proteins such as titin, whose native structures are micrometers long (8). In this chapter, we present a method that extends the use of optical tweezers to study the unfolding and refolding trajectories of individual globular proteins (9).

This method relies on the use of molecular handles, ∼500 bp DNA molecules, to connect the protein to polystyrene beads and minimize the interactions between the tethering surfaces of the bead (see Fig. 1). One end of each DNA molecule is covalently attached to a cysteine residue of the protein through a disulfide bond, while the other end is bound to a bead through either streptavidin–biotin or digoxigenin–antibody interactions. During the experiment, the protein is stretched and relaxed by moving the tethered beads relative to each other by means of piezoelectric actuators. We chose DNA molecules as our molecular handles because they are easy to synthesize in the laboratory by polymerase chain reaction (PCR) and because their mechanical ­properties have been extensively characterized by optical tweezers (10, 11), making it possible to distinguish their contributions easily in the recorded traces. In this chapter, we detail the protocols to generate DNA–protein chimeras and manipulate them with optical tweezers. We primarily consider DNA molecules made of ∼500 bp because they have been the handles of choice for most of our experiments; handles of different lengths, however, can be used (9).
Fig. 1.

Experimental setup (not to scale). A single protein is manipulated between two micrometer-sized polystyrene beads by means of DNA molecular handles. One handle binds to a bead (held in the optical trap) through a digoxigenin–antibody interaction; the other handle binds to a bead (held by suction to the end of a pipette) through a streptavidine–biotin interaction. Each handle is covalently bound to the protein through a disulfide bond (see Fig.  2). During the experiment, the protein tethered between the beads is stretched and relaxed by moving the pipette relative to the optical trap by means of a piezoelectric actuator. (Adapted from Fig.  3 of ref. 13 with kind permission from IOS Press).

The method described in this chapter represents a unique approach to single-molecule manipulation studies and presents a number of unique capabilities. (1) Individual globular proteins are directly manipulated with no need for the engineering of polymeric proteins, such as AFM experiments require. (2) The DNA handles are attached to cysteine residues that can be located anywhere on the surface of the protein, allowing a multitude of different pulling geometries to be used to study the anisotropy of the molecule’s energy landscape. (3) The attachment of the protein–DNA chimeras to the beads is achieved through specific bonds (biotin–streptavidin or digoxigenin–antibody), ensuring reproducibility between different experiments and easy interpretation of the data. (4) Unwanted multiple attachments between the tethering surfaces can be easily recognized. The response of DNA to force is known to be characterized by an overstretching transition at approximately 65 pN (11). If two or more molecules are caught between the two beads and pulled in parallel, the DNA molecules share the load and overstretch at higher forces. (5) The low spring constant of the optical trap and the force resolution of the instrument (RMS force noise of ∼0.5 pN) allow the refolding process – as well as fluctuations between different molecular conformations – to be monitored directly.

This method has been used to manipulate RNase H (12, 13) and T4 lysozyme (14) and should be applicable to any protein in which two exposed cysteine residues can be engineered. The DNA handles do not seem to affect the stability and folding mechanism of the tethered proteins, as shown by circular dichroism studies (9). Moreover, RNase H bound to two 558 bp molecular handles retains its enzymatic activity, proving that the overall three-dimensional structure of the tethered protein is conserved (12).

This method presents some drawbacks as the components required to generate the DNA–protein chimeras are quite ­expensive and the range of forces that can be applied is limited to between 0 and 64 pN; at higher forces, the DNA overstretches and the digoxigenin/antibodies interaction becomes very labile, making interpretation of the data difficult. It should be noted, however, that the probability of observing a protein unfold depends not only on the applied force but also on the loading rate (r) used to pull on the molecule; r (pN/nm) equals pulling speed (nm/s) times spring constant (pN/nm). Owing to the low spring constant of an optical trap, the loading rates used in optical tweezer experiments are quite low and thus even proteins that are mechanically quite resistant can be observed to unfold below 64 pN of force.

2 Materials

2.1 Generation of Protein Variants

  1. 1.

    QuikChange site-directed mutagenesis kit (Stratagene).

     
  2. 2.

    High copy plasmid under a T7 promoter, such as pAED4 (from the Paul Matsudaira lab, The Whitehead Institute, Cambridge, MA, USA) or pET27 (Novagen).

     
  3. 3.

    Expression strain of Escherichia coli cells (BL21(DE3)plysS) (Promega).

     
  4. 4.

    Dithiothreitol (DTT) or Tris(2-carboxyethyl)phosphine (TCEP).

     
  5. 5.

    Column equilibration buffer: 0.1 M sodium phosphate buffer, pH 5.5.

     
  6. 6.

    Column equilibration buffer: 0.1 M sodium phosphate buffer, pH 7.0.

     
  7. 7.

    Disposable 5 mL polypropylene columns (Pierce).

     
  8. 8.

    Sodium azide (NaN3).

     
  9. 9.

    Sephadex G-25 coarse resin (Sigma).

     
  10. 10.

    DTPD solution: a 10 mM 2,2′-dithiodipyridine solution is prepared by first dissolving 22.22 mg of DTDP in 1.5 mL of acetonitrile and then adding pH 5.5 column equilibration buffer to a final volume of 10 mL. Aliquots of 22.22 mg of DTDP dissolved in 1.5 mL of acetonitrile can be prepared and stored at −80 °C for months.

     
  11. 11.

    pH 5.5/G-25 columns: three disposable 5 mL polypropylene columns are cast with Sephadex G-25 coarse resin previously swelled overnight (O/N) in 0.2% NaN3 at room temperature (RT). The gel is allowed to settle in the column for at least 30 min at RT and it is then equilibrated with five column volumes of pH 5.5 column equilibration buffer, flown through via gravity. Immediately before being used for buffer exchange (i.e., loaded with protein), these columns must be spun “dry” at 1,000  ×  g for 1.5 min to remove excess buffer (seeNote 1).

     
  12. 12.

    pH 7.0/G-25 columns: three disposable 5 mL polypropylene columns are cast with Sephadex G-25 coarse resin previously swelled overnight (O/N) in 0.2% NaN3 at RT. The gel is allowed to settle in the column for at least 30 min at RT and it is then equilibrated with five column volumes of pH 7.0 column equilibration buffer, flown through via gravity. Immediately before being used for buffer exchange (i.e., loaded with protein), these columns must be spun “dry” at 1,000  ×  g for 1.5 min to remove excess buffer.

     
  13. 13.

    Ready Gel® precast polyacrylamide gels 4–20% (Bio-Rad).

     

2.2 Generation of DNA Molecular Handles

  1. 1.

    PTC-200 Peltier thermal cycler (MJ Research).

     
  2. 2.

    Taq DNA polymerase (Qiagen).

     
  3. 3.

    pGEMEX1 plasmid DNA (Promega).

     
  4. 4.

    DNA primers (Integrated DNA Technology).

     
  5. 5.

    HiSpeed Plasmid Maxi kit (Qiagen).

     
  6. 6.

    Maxi kit DTT column elution buffer: 15 mM NaPO4 and 3 mM DTT, pH 7.0.

     
  7. 7.

    Handle buffer: 15 mM NaPO4 and 3 mM DTT, pH 7.0.

     
  8. 8.

    30 MWCO Microcon ultrafiltration cartridges (Millipore).

     
  9. 9.

    Micro Bio-Spin columns with Bio-Gel P-6 in Tris buffer (Bio-Rad).

     
  10. 10.

    Spin column buffer: 0.1 M NaPO4 and 1 mM EDTA, pH 8.0.

     

2.3 DNA–Protein Coupling

  1. 1.

    PAGE reagents: 30% acrylamide/bisacrylamide 29:1, Tris(hydroxymethyl)aminomethane hydrochloride, sodium dodecyl sulfate, ammonium persulfate, and N,N,N′,N′-tetramethylethylenediamine.

     
  2. 2.

    Ready Gel® precast polyacrylamide gels 4–20% (Bio-Rad).

     
  3. 3.

    SYBR Green I nucleic acid gel stain (Molecular Probes). SYPRO Red protein gel stains (Molecular Probes). Typhoon 8600 (Molecular Dynamic). DNA silver stain kit (GE Healthcare).

     
  4. 4.

    AFM deposition buffer: 10 mM HEPES, 10 mM NaCl, and 2 mM MgCl2 (pH 7.5).

     
  5. 5.

    A Veeco Nanoscope III AFM was used by us.

     
  6. 6.

    Pointprobes, type NCH-100 AFM probes (Nanosensors).

     
  7. 7.

    Ruby mica sheets (Asheville–Schoonmaker).

     

2.4 Preparation of Anti-digoxigenin Antibody-Coated Beads

  1. 1.

    PBS (7.0): 0.14 M NaCl, 2.7 mM KCl, 61 mM K2HPO4, and 39 mM KH2PO4, pH adjusted to 7.0 with HCl.

     
  2. 2.

    PBS (7.4): 0.14 M NaCl, 2.7 mM KCl, 80.2 mM K2HPO4, and 20 mM KH2PO4, pH adjusted to 7.4 with HCl.

     
  3. 3.

    Cross-linking buffer: 100 mM Na2HPO4, pH 8.5, and 100 mM NaCl (or other non-amine containing buffer pH 7–9).

     
  4. 4.

    Protein G-coated polystyrene particles, 3.18 μm diameter, 0.5% w/v, 5 mL (Gentaur Molecular Products).

     
  5. 5.

    Anti-Dig solution: 1 mg/mL anti-digoxigenin (anti-Dig) antibody solution prepared by dissolving 200 μg of sheep polyclonal anti-Dig antibody (Roche) in 200 μL of PBS (pH 7.4).

     
  6. 6.

    DMP solution: A 10 mM dimethyl pimelimidate (DMP) (Pierce) solution is prepared by dissolving 50 mg of DMP in 1 mL of cross-linking buffer. DMP must be used immediately after its preparation because of its instability.

     
  7. 7.

    Tris base solution: 2 M Tris.

     

2.5 Tethering of Protein–DNA Chimeras to Polystyrene Beads

  1. 1.

    Binding buffer:10 mM Tris, 250 mM NaCl, and 10 mM MgCl2, pH 7.0.

     
  2. 2.

    Streptavidin-coated, 2.10-μm beads (Spherotech).

     

3 Methods

Each molecular handle is attached to a protein via the formation of a disulfide bond between a thiol group present at the end of the DNA molecule and a thiol group of a cysteine residue in the ­protein (Fig. 1). The thiol–thiol reaction is mediated by DTDP (15, 16, 17) (Fig. 2). Disulfide bonds can form spontaneously, especially at high pH and temperatures (18, 19). However, DTDP speeds up the reaction and allows the time course of the DNA–protein coupling to be monitored spectrophotometrically at 343 nm via the release of the leaving group pyridine-2-thione (9).
Fig. 2.

Schematic of the reactions used to attach DNA molecules covalently to proteins. (a) A cysteine-bearing protein is first activated with DTDP. (b) After cleaning up the reaction product, it is then allowed to react with DNA molecules bearing a thiol group at one end. The kinetics of both the protein thiol–pyridine activation and the protein–DNA coupling can be followed spectrophotometrically at 343 nm via the release of the leaving group pyridine-2-thione. (Adapted with kind permission of Springer Science  +  Business Media from ref. 9).

DTDP can be used to activate thiol groups of either DNA or proteins; in our experiments, however, we activate proteins because in this way the success of the reaction can be assessed by mass spectroscopy and because the activated molecule can then be used to generate polymeric proteins if desired. In the following sections, we describe in detail protocols to generate DNA–protein chimeras and to use them in single-molecule optical tweezer experiments.

3.1 Generation and Activation of Cysteine-Bearing Protein Mutants

The following is the method to activate monomeric proteins for coupling with DNA, and the variant for coupling protein oligomers. Some proteins spontaneously react to form polymeric ­proteins without DTDP if stored in the absence of a reducing agent (18). However, in our experience, the speed of the reaction is greatly accelerated and more likely to continue to completion when DTDP is used.

Generation of Monomeric Proteins

  1. 1.

    A protein variant bearing only two cysteine residues exposed on the surface of the protein (seeNote 2) is generated through site-directed mutagenesis. Other cysteines naturally occurring in the protein are substituted with structurally neutral residues, such as alanine.

     
  2. 2.

    The cysteine-bearing variant is purified according to the protocol used for its cysteine-free variant with the only difference being that 1 mM DTT is added to all purification solutions (seeNote 3). Depending on the inherent stability of the protein, the purified protein can be kept soluble at 4 °C for short-term storage, or at −80°C after either lyophilization or liquid nitrogen freezing for long-term storage.

     
  3. 3.

    Before DTDP activation, the purified cysteine-bearing protein in buffer at pH 7.0 is allowed to react with 10–30 molar excess of DTT or 10 mM final TCEP (seeNote 3) for ∼1 h at RT to ensure full reduction of the thiol groups.

     
  4. 4.

    A volume of 150–200 μL (300 μL max) of reduced protein is carefully loaded onto one pH 5.5/G-25 column placed in a 15 mL conical tube and spun down at 1,000  ×  g for 1.5 min; during loading, attention should be paid to avoid disrupting the resin bed. To prevent the formation of protein oligomers after removal of the reducing agent, 150–200 μL of 10 mM DTDP is placed at the bottom of the 15 mL Falcon tube prior to centrifugation.

     
  5. 5.

    The initial reaction of DTDP with a new protein variant should be monitored spectroscopically at 343 nm for production of the leaving group pyridine-2-thione (seeNote 4 and Fig. 2). The activation of the cysteine residues is usually complete in a few minutes as seen by the time course of the release of the leaving group pyridine-2-thione during the reaction (9), although less accessible cysteines might take longer. Even rapid reactions that appeared to be complete in a matter of minutes were typically permitted to continue O/N at RT to ensure complete activation of the protein by DTDP. Some research groups, however, may elect to accelerate sample preparation for very reactive protein variants by allowing the reaction to proceed only for 2–4 h.

     
  6. 6.

    The thiol–pyridine-activated protein is isolated from the excess of unreacted DTDP by sequentially spinning the reaction solution through two pH 5.5/G-25 columns at 1,000  ×  g for 1.5 min. The protein is now activated and ready to be attached to DNA handles.

     
  7. 7.

    The activated protein can be stored at 4 °C for several days (depending on the inherent stability of the protein).

     
  8. 8.

    The success of the thiol–pyridine activation can be assessed through mass spectroscopy (9). This is particularly recommended for the initial reactions with new protein substrates to ensure complete activation by DTDP.

     

Synthesis of Polymeric Proteins

  1. 1.

    Reducing agents were removed from nonactivated protein by sequential spins through two pH 7.0/G-25 columns at 1,000  ×  g for 1.5 min (seestep 3 above and Note 3). Because polymers are desired in this case, the nonactivated protein does not need to be reduced with DTT or TCEP as was done prior to DTDP activation.

     
  2. 2.
    Thiol–pyridine-activated protein (from step 8 above) in pH 5.5 column equilibration buffer is mixed with nonactivated protein at a molar ratio of 1:2 (see Fig. 3a). Typically, the final concentration of the activated protein is ∼25 μM. The reaction is allowed to proceed O/N at RT.
    Fig. 3.

    Synthesis of polymeric proteins. (a) Thiol–pyridine-activated proteins are mixed with thiol-modified proteins to generate tandem repeats of the molecule (adapted from ref. 9, with kind permission of Springer Science+Business Media). (b) SDS–PAGE analysis of an RNase H*Q4C/V155C polymerization reaction: unreacted protein (lane 1) and the product of the reaction (lane 2  ). As expected, the product of the reaction is a population of polymers of different lengths. The gel was stained with SYPRO Red, analyzed with a Typhoon 8600, and is shown in gray scale. (c) Intensity profile of the gel along the dark line in lane 2. Each peak of the profile corresponds to a different polymerization product, starting with the monomer from the left. The longest detectable polymers for this reaction were made of 24 monomers.

     
  3. 3.

    The following day, more thiol–pyridine-activated protein (typically ¼ of the amount used in the first reaction) is added to the polymerization solution and allowed to again react O/N at RT. This second step aims to cap the ends of the polymers with activated monomers, which can then react with molecular DNA handles bearing thiol groups.

     
  4. 4.

    The degree of polymerization reached can be assessed using a SDS–PAGE gradient gel (4–20%) (see Fig. 3b, c).

     

3.2 Preparing DNA Molecular Handles

  1. 1.

    Handles of 558 bp were generated by PCR. One handle was synthesized using the primers 5′-thiol-GCT-ACC-GTA-ATT-GAG-ACC-AC and 5′-biotin-CAA-AAA-ACC-CCT-CAA-GAC-CC. The other handle was synthesized using the same 5′-thiol primer together with 5′-digoxigenin-CAA-AAA-ACC-CCT-CAA-GAC-CC. PCR was performed using Taq polymerase and pGEMEX1 as a template. DTT at a final concentration of 20 mM was added to the PCR to keep the thiol groups of the primers reduced. Large amounts of handles (400–500 μg) were typically synthesized using as much as 9 mL of PCR reactions (PCR conditions are optimized for production yield and purity). The DNA handles were then purified using HiSpeed plasmid maxi kit columns and eluted with maxi kit DTT column elution buffer. The purified handles can be stored at −20 °C for months.

     
  2. 2.

    The two types of handles are mixed in equal amounts to obtain ∼1 mL of ∼200 μg/mL digoxigenin/biotin (dig/bio) handles in handle buffer.

     
  3. 3.

    The dig/bio handle solution is then concentrated down to 50–60 μL with a 30-kDa MWCO Microcon centrifuge tube.

     
  4. 4.

    Reducing agents are removed from the handles by sequentially spinning them through three Micro Bio-Spin P6 columns equilibrated with the spin column buffer. The final handle concentration is usually 20–25 μM.

     

3.3 DNA–Protein Coupling

Monomeric Proteins

  1. 1.

    The dig/bio handles are mixed with thiol–pyridine-activated protein in pH 5.5 column elution buffer in a DNA handles:protein molar ratio of 4:1; typically, ∼20 μM of DNA handles are reacted with ∼5 μM of activated protein. The reaction is allowed to proceed O/N at RT (seeNote 5).

     
  2. 2.
    The kinetics of the attachment of DNA molecules to proteins is slow, usually taking between 24 and 48 h to reach completion (9). The extent of the DNA–protein coupling can be assessed either by a 4% SDS–PAGE gel, prepared according to ref. 20, or by AFM (see Fig. 4).
    Fig. 4.

    Characterization of the protein–DNA coupling reaction. (a) AFM image of the product of the reaction between ­558 bp DNA molecules and RNase H*Q4C/V155C. Two protein–DNA chimeras made of two handles and one RNase H molecule are clearly visible. (b) 4% SDS–PAGE of the attachment of DNA handles to T4 lysozyme*T21C/K124C: DNA handle alone (lane 1), proteins bound to one handle (lane 2  ), handle dimers formed through the formation of disulfide bonds between the DNA thiol groups (lane 3  ), and protein bound to two handles (lane 4  ). The gel was stained with SYBR Green II, analyzed with a Typhoon 8600, and is shown in gray scale. (Adapted from ref. 9, with kind permission of Springer Science  +  Business Media)

     
  3. 3.

    To visualize the product of the protein–DNA reaction by AFM, molecular constructs were diluted to a final concentration of 2 nM in AFM deposition buffer. A volume of 20 μL of solution was deposited onto freshly cleaved mica and allowed to adsorb onto the surface for 1 min. The mica ­surface was then gently washed with doubly distilled water and dried with a stream of nitrogen. The sample was imaged in air in tapping mode.

     

Polymeric Proteins

  1. 1.
    After the polymerization reaction, the longer polymers can be isolated from smaller protein oligomers by gel filtration (see Fig. 5). This step, although not strictly necessary, ensures that only long polymers are used for single molecules studies, which provides more data per single pull of the polymeric sample.
    Fig. 5.

    Isolation of long polymeric proteins through gel filtration. A 4–20% gradient SDS–PAGE gel of polymers of RNase H*Q4C/V155C (lane 1) and of different gel filtration fractions, starting from the latest (lane 2  ) and finishing with the slowest (lane 6  ). Gel filtration was performed using a size-exclusion column TosoHaas G2000 SWXL, equilibrated with 300 mM NaCl and 25 mM NaPO4, pH 7.4, and run at 0.75 mL/min. The gel was stained with SYPRO Red and analyzed with a Typhoon 8600, and is shown in gray scale. Each fraction was collected manually and they were each approximately 1 mL in volume.

     
  2. 2.

    The polymeric proteins are then coupled to DNA handles following the same protocol used with monomers. For this reaction, the concentration of polymeric protein ends can be estimated from the overall absorbance of the protein sample and from the intensity of the different bands in polyacrilamide gels. Depending on the relative sizes of the DNA handles and proteins being used, the success of the reaction can be assessed either by SDS–PAGE (9) or simply by manipulating the molecules using optical tweezers (see Subheading 3.4).

     

3.4 Preparation of Anti-digoxigenin Antibody-Coated Beads

  1. 1.

    Protein G-coated polystyrene beads are spun down on a bench centrifuge at 1,000  ×  g for 5 min and then resuspended in 1 mL of cross-linking buffer.

     
  2. 2.

    A volume of 60 μL of anti-Dig solution and 30 μL of DMP solution are added to the resuspended protein G-coated beads and the reaction is tumbled at RT for 60 min. The beads are then spun down at 1,000  ×  g for 5 min, resuspended in 1 mL of Tris base solution, and vortexed for 2 h at RT at minimal speed to quench the reaction. Notice that the Tris base quenching is a recommended but not necessary step. The beads are then diluted threefold in PBS (7.0), spun down at 1,000  ×  g for 15 min, and resuspended in PBS (7.0) three times to isolate the beads from the unreacted antibodies and DMP molecules. The beads are now ready to be used in laser tweezer experiments and can be kept at 4 °C for months.

     

3.5 Tethering of Protein–DNA Chimeras to Polystyrene Beads

  1. 1.
    Protein–DNA chimeras are allowed to react with anti-Dig beads in binding buffer for about 15 min at RT. The volume of a typical reaction was 10 μL total, with 8 μL of buffer, 1 μL of beads, and 1 μL of diluted DNA–protein chimera (3–20 pM final protein concentration in the 10 μL reaction). At the end of the reaction, the DNA–protein chimeras will be attached to the beads via the dig-labeled handle, while the other bio-labeled handle remains free and available for binding to streptavidin (see Fig. 6).
    Fig. 6.

    Schematic of the experimental procedure used to tether single proteins in ­optical tweezer experiments. (a) Polystyrene beads covered with antibodies against digoxigenin are first allowed to react with the DNA–protein chimeras for 15 min at RT. (b) The DNA–protein chimera beads are then flown into the fluid chamber of the optical tweezer instrument and caught in the optical trap. A second bead covered with streptavidin was previously attached to a micropipette by suction. This streptavidin bead is then brought close to the optical trap bead to facilitate its binding to the DNA biotin moiety on the free end of the DNA–protein chimera.

     
  2. 2.

    A streptavidin bead was suctioned onto the micropipette tip in the optical tweezer chamber.

     
  3. 3.

    The anti-Dig beads – now bearing protein–DNA chimeras – are further diluted into binding buffer (∼1 mL) and then flowed into the optical tweezer fluid chamber until one bead is caught. The micropipette streptavidin bead is brought into close proximity to the anti-Dig chimera bead to allow the DNA–biotin moiety to react with streptavidin. Once the biotin/streptavidin bond is formed, the protein–DNA chimera is tethered between the two differently derivatized beads.

     

3.6 Manipulation of Protein–DNA Chimeras with Optical Tweezers

Monomeric Proteins

Once tethered between two beads, a protein–DNA chimera can be stretched and relaxed multiple times by moving the pipette relative to the optical trap. In our setup, the applied force is determined by measuring the change in light momentum of the beams leaving the optical trap, while the extension of the molecule is determined by means of a “light lever system” (21). Under tension, the unfolding of a protein is associated with a large change in its extension, as the molecule goes from a compact native state to an elongated unfolded state; on the contrary, the refolding process causes a sharp compaction of the protein. These sudden changes in molecular end-to-end distance give rise to sharp transitions in the force vs. extension traces.

As an example, Fig. 7a shows a force–extension curve obtained by manipulating an RNase H molecule. By analyzing the size of the transitions in the recorded traces, it is possible to calculate the number of amino acids involved in the unfolding or refolding processes. Due to the low spring constant of the optical trap (usually 0.1 pN/nm or less), fluctuations between different molecular conformations can be monitored in real time. Figure 7b shows an extension vs. time trace obtained by keeping an RNase H molecule at a constant force using the force-feedback mode of the instrument; under these experimental conditions, the protein is observed to fluctuate between its intermediate and unfolded states.
Fig. 7.

Mechanical manipulation of single protein by optical tweezers. (a) Force–extension curve obtained by stretching and relaxing an RNase H*Q4C/V155C molecule. The protein unfolds at about 17 pN (native state N  →  U unfolded state) and refolds into an intermediate state (I  ) at about 5 pN (U  →  I  ). The intermediate state refolds into the native structure at lower forces ( I  →  N). (b) Extension vs. time trace showing an RNase H*Q4C/V155C protein hopping between its unfolded and intermediate states when holding the protein at a constant force of 5.5 pN. The extension values are the distances between the centers of the tethering beads.

Polymeric Proteins

When a polymeric protein is stretched and relaxed, the subsequent unfolding and refolding of the individual domains give rise to characteristic saw-tooth patterns in the force–extension traces (Fig. 8). Each tooth corresponds to the unfolding or refolding of an individual monomer within the polymeric protein–DNA chimera. The number of teeth in the force–extension traces will vary because of the heterogeneity in length of the polymeric proteins generated through the method described above; typically, however, between four and ten teeth were observed in our curves. Pulling on polymers presents the advantage of providing information on the mechanical properties of several domains at the same time, and thus it is easier to collect a statistically relevant set of data. The interpretation of the results, however, is usually more difficult than with monomeric proteins.
Fig. 8.

Mechanical manipulation of a polymeric protein. Force–extension cycles obtained by stretching and relaxing a polymer of RNase H*Q4C/V155C multiple times. (Reprinted with kind permission of Springer Science  +  Business Media from ref. 9).

4 Notes

  1. 1.

    We use pH 5.5 for protein activation by DTDP to keep the reaction conditions stringent for activation of cysteine residues. However, at higher pH, the rate of this reaction and the rate of protein polymerization increases, and this faster kinetics could be useful with protein variants that react more slowly than those we investigated. Similar buffer exchange columns at pH 7.0 will be used to prepare other reaction components later in the protocols; they will be referred to as “pH 7.0/G-25 columns.”

     
  2. 2.

    It is technically possible to construct protein variants for optical tweezer experiments in which the structure of the protein is unknown or the engineered cysteines are not on the exposed protein surface. For instance, denaturants can be used to expose buried cysteines transiently and permit DNA handle attachment, but the disrupted structures of these DNA–protein chimeras will prevent any meaningful data from being obtained.

     
  3. 3.

    DTT is preferred over β-mercaptoethanol during protein purification because it is a more efficient reducing agent. TCEP, a very potent reducing agent, can also be used. We recommend TCEP for storing concentrated, pure proteins and for ensuring complete reduction of cysteines prior to activation with DTDP, but it is too expensive to justify its use in protein purification buffers.

     
  4. 4.

    The activation rate of a particular protein variant with DTDP varies depending on the accessibility of the cysteine, necessitating spectroscopic monitoring during the initial reaction. We also recommend monitoring the ratio of the 343 and 405 nm absorbance values rather 343 nm alone to eliminate any potential shifts in the baseline due to either instrument drift or protein precipitation. Some proteins did exhibit minor precipitation that caused an anomalous baseline shift; this precipitation appeared to be variant specific and potentially related to the final concentration of acetonitrile (used to dissolve DTDP) in the reaction. Proteins that are more sensitive to the presence of organic solvents could be accommodated by changing the volumetric ratios of the DTDP and the ­protein stock solutions, while preserving the molar ratios, to decrease the final concentration of acetonitrile in the reaction.

     
  5. 5.

    Only 50% of the protein–DNA chimeras synthesized through this method bear the correct combination of handles: that is, one handle labeled with biotin and the other with digoxigenin. The rest of the chimeras carry identically labeled handles. However, because such chimeras cannot be tethered between two differently derivatized polystyrene beads, these molecules are not functional in optical tweezer experiments. The use of these mixed populations of DNA–protein chimeras – rather than a pure population of sample containing one digoxigenin- and one biotin-labeled DNA handle – also does not appear to impact the optical tweezer experiments negatively: identical data were obtained from mixed sample populations and those engineered to contain a pure population (9).

     

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Copyright information

© Springer Science+Business Media, LLC 2011

Authors and Affiliations

  • Ciro Cecconi
    • 1
  • Elizabeth A. Shank
  • Susan Marqusee
  • Carlos Bustamante
  1. 1.CNR-Istituto Nanoscienze S3, Department of PhysicsUniversity of Modena e Reggio EmiliaModenaItaly

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