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Recent advances in optical imaging and molecular manipulation techniques have made it possible to observe the activity of individual enzymes and study the dynamic properties of processes that are challenging to elucidate using ensemble-averaging techniques. The use of single-molecule approaches has proven to be particularly successful in the study of the dynamic interactions between the components at the replication fork. In this section, we describe the methods necessary for in vitro single-molecule studies of prokaryotic replication systems. Through these experiments, accurate information can be obtained on the rates and processivities of DNA unwinding and polymerization. The ability to monitor in real time the progress of a single replication fork allows for the detection of short-lived, intermediate states that would be difficult to visualize in bulk-phase assays.
DNA replication involves the coordinated activity of a large number of proteins. The replisome, the molecular machinery of DNA replication, unwinds the double-stranded DNA, provides primers to initiate synthesis, and polymerizes nucleotides onto each of the two growing strands (1). Remarkable progress has been made in characterizing the structural and functional properties of the individual components; their coordination at the replication fork is less well understood.
The dynamic nature of the replisome makes it hard to probe its coordination with ensemble-averaging techniques. We describe here single-molecule techniques to observe, in real time, the replication of individual DNA molecules by replication complexes of E. coli and the bacteriophage T7. Replication reactions from both these systems can be reconstituted in vitro with a relatively small number of proteins and are compatible with the optical imaging and manipulation techniques developed to study DNA-protein interactions at the single-molecule level (2–4).
In this chapter, we describe how individual DNA molecules can be mechanically stretched and their lengths used as a probe for enzymatic activity at the replication fork. Linearized λ DNA is modified to have a biotin on one end and a digoxigenin moiety on the other. The biotinylated end is attached to a functionalized glass coverslip and the digoxigeninated end to a small bead (Figure 1). The assembly of these DNA-bead tethers on the surface of a flow cell allows a laminar flow to be applied and a drag force on the bead to be exerted. As a result, the DNA is stretched close to and parallel to the surface of the coverslip at a force that is determined by the flow rate. The length of the DNA is measured by monitoring the position of the bead. Length differences between single- and double-stranded DNA are utilized to obtain real-time information on the activity of the replication proteins at the fork (2–4) (Figure 2).
To minimize nonspecific interactions between the glass surface and proteins, we covalently couple high-molecular-weight polyethylene glycol (PEG) to the surface. First, the glass is coupled to the alkoxy group of an aminosilane, creating a surface with reactive amine groups that can subsequently be coated with a polymer of choice (5). Here, we describe how a mixed population of biotinylated and non-biotinylated succinimidyl propionate-PEG is coupled to the amine-functionalized glass, coating the coverslip in a layer of PEG displaying a mixture of biotin and nonreactive methyl groups. The biotin is used to tightly bind streptavidin, allowing for a subsequent coupling of biotinylated DNA to the surface. Any functionalized PEG can be used to allow a customizable surface based on choice of DNA modification.
Bacteriophage λ DNA is 48.5 kb of double-stranded DNA readily purchased from suppliers, providing an ideal scaffold for single-molecule DNA manipulation. The linearized DNA has 12-base single-stranded overhangs at each end, to which we attach modified and unmodified oligonucleotides using standard annealing and ligating techniques. The following steps describe in detail how to prepare a DNA substrate with a primed replication fork at the surface-attached end and a site for bead attachment at the other (Figure 1). The following protocol will result in 0.5 mL of DNA substrate at a concentration of 1.4 nM.
In the flow-stretching single-molecule experiment, we measure DNA length change by observation of the position of a small bead bound to the end of the λ DNA. To achieve this, the beads are functionalized with a Fab fragment with specificity for the digoxigenin. Activated beads can then be attached to tethered DNA and used to manipulate the DNA. The following protocol will result in 1.0 mL of 1–2×109 beads/mL functionalized beads.
Once the DNA and functionalized beads have been prepared and microscope coverslips have been functionalized, a flow cell can be assembled and single-molecule experiments performed. Here we describe how a flow chamber is prepared with the functionalized coverslip and a quartz slide, and how the substrate is constructed in situ by flowing λ DNA fork substrates and functionalized beads.
Solutions containing replication proteins are introduced into the flow cell and any bead movement is observed in real time by imaging the tethered bead positions with a CCD camera. Movement of the beads is converted to length change of the DNA, allowing temporal and kinetic analysis of single replication events. Several methods to relate DNA length changes to replication can be employed. In the first method, we make use of the fact that at stretching forces lower than 6 pN, DNA in the single-stranded form is considerably shorter than DNA in the duplex form (Figure 1d). In the case of leading-strand synthesis, only one of the two unwound parental DNA strands will be converted to duplex DNA. In the absence of lagging-strand synthesis, the other strand will remain in the single-stranded form after helicase-mediated unwinding at the fork. By attaching the 5′ lagging strand of the DNA to the surface, we can observe leading-strand synthesis by the effective conversion of parental duplex DNA into single-stranded DNA, resulting in a shortening of the DNA. The second method is employed in those reactions with both leading- and lagging-strand synthesis. Here, no net conversion between single- and double-stranded DNA takes place, but the transient formation of a replication loop at the lagging strand can be observed as a brief and gradual shortening of the DNA, followed by a sudden lengthening. These two events correspond to the formation and release of a replication loop, respectively.
The authors would like to thank Charles Richardson and Nick Dixon for their generous gifts of T7 and E. coli replication proteins, respectively. The authors would also like to acknowledge contributions from Paul Blainey, Candice Etson, Jong-Bong Lee, and Joseph Loparo towards the development of the single-molecule replication assay.
1When used in a single-molecule experiment, observation of tethered λ DNA serves as an internal control that the substrate is assembled properly. As the biotinylated oligo does not complement the λ itself but rather the other fork arm, if DNA is attached to the surface it immediately shows correct fork assembly. Any bead attached similarly confirms annealing of the digoxigenin oligo. As a caveat, the fork oligos are present at a high concentration and can anneal without the λ DNA to form small forks which can bind to the surface. This is typically not a matter of concern, but for troubleshooting or adapting the technique to higher resolution experiments or other protein systems this fact should be considered. Eliminating the excess forks is simply a matter of reducing the oligonucleotide ratios in substrate construction or purifying the free oligos away after preparation of the DNA constructs.
2The PEG powders and silane solutions are extremely sensitive to hydrolysis, and care must be taken to prevent their degradation. Immediately after use, place containers in a desiccator and remove air. Replace with inert atmosphere (N2 or Ar) and seal lid with plastic wrap until next use. Batches of functionalized coverslips that display poor tethering capability are frequently due to degraded reagents.
3Syringe pumps often exhibit small irregularities in flow, resulting in significant force fluctuations. A simple way to reduce these flow instabilities is to place an airspring between the flow cell and the syringe pump (see Figure 1b). A 50 ml plastic tube is sealed and the lid affixed using epoxy. Three holes are pierced in the lid, and three lengths of tubing (same tubing as flow cell) are inserted to ~1 cm from the bottom. Using epoxy, the tubes are sealed to the lid, preventing any air from entering or escaping. The tube is filled with 40–45 mL water and connected to the syringe pump with one of the three tubing pieces. The remaining two connect to the flow cell tubing using an adaptor piece of slightly larger tubing. Upon starting the syringe pump, the withdrawal of water from the air spring will result in a pressure drop in the closed air volume. This negative pressure will cause buffer to flow through the flow cell. Any irregularity in the syringe pump will not immediately change the negative pressure in the air spring and will be dampened out very effectively. The airspring provides two additional benefits: 1) a simple method of connecting the two outlet channels to a single syringe pump, and 2) an easy way of changing the flow direction. By lifting the airspring, gravity will force the flow to reverse direction and cause the bead-DNA tethers to flip back and forth with the flow.
4A common problem in the experiment occurs when the large beads nonspecifically stick to the surface of the coverslip, preventing movement and measurement. As a solution, we apply a small magnetic force (~1.7 pN) perpendicular to the flow direction to lift the beads off the surface. Permanent rare-earth magnets are moved into place above the flow cell using a 2-axis translational stage (Thor Labs) immediately prior to data acquisition.
5Dark-field illumination can be used to increase the contrast in the bead imaging. A fiber illuminator (Thor Labs OSL1) is positioned at an incidence angle between 10 degrees and parallel to the microscope stage ~0.5m away. The low numerical aperture of the 10X objective will not allow the illumination light to be transmitted, but will allow the light scattered by the beads to be imaged. As a result, the beads can now be seen as bright objects against a dark background.