Analysis of metal-dependent DNA nicking activities by Cas endonucleases

Abstract

CRISPR-Cas systems use RNA-guided CRISPR-associated (Cas) effectors to neutralize infections in bacteria and archaea. In class 2 CRISPR-Cas systems, Cas9 and Cas12 are single-protein Cas effectors that target double-stranded DNA based on complementarity to the guide RNA before cleaving the target DNA using metal-dependent endonuclease domains. Cas9 and Cas12 proteins can be readily programmed to target any DNA of interest by changing the guiding RNA sequence and have been co-opted for genome editing and other biotechnology purposes. The effect of metal ion concentration is an essential consideration in the physiological role of Cas immunity effectors as well as the biotechnological applications of Cas endonucleases. In this chapter, we describe methods for studying the effect of variable divalent metal ion conditions on the DNA binding and cleavage activities of well-studied Cas9 and Cas12a proteins.

1. Introduction

The effectors of CRISPR-Cas immune systems are Cas proteins guided by CRISPR RNAs (crRNAs) to bind to complementary nucleic acids [1]. Cas9 and Cas12a are popular effector proteins that target DNA [2-4]. Many orthologs of each protein have been extensively researched and several are effective tools for genome editing [4-11]. DNA binding by Cas9 and Cas12a is initiated by searching for protospacer adjacent motif (PAM) sequences [12-16]. PAMs are short and conserved motifs located next to target “protospacers” [17], and are crucial for target location and DNA unwinding [14,18,19]. At PAM sites, Cas9 and Cas12a endonucleases momentarily pause to assess if the adjacent sequence matches the guiding crRNA “spacer” sequence (Fig. 1) [12,16]. If a complementary sequence is found, the Cas endonuclease triggers DNA unwinding by forming crRNA-DNA base pairs, starting at the PAM-proximal region of the protospacer. This unwinding progresses directionally away from the PAM to form an R-loop [20]. Consequently, the initial base pairs formed in the PAM-proximal, or “seed”, region play a more significant role in target binding than those in the PAM-distal region [2,4].

Figure 1: DNA binding and cleavage mechanisms for (A) Cas9 and (B) Cas12a effectors.

Both Cas effectors search for targets and initiate DNA unwinding based on the presence of a PAM sequence adjacent to the target. DNA unwinding occurs directionally away from the PAM. The overall process of DNA binding is dependent on Mg²⁺. (A) Following complete formation of the R-loop, the Cas9 HNH domain undergoes a Mg²⁺-dependent conformational rearrangement that is necessary for cleavage of the target strand. Both the HNH and the RuvC domain require divalent metal ion cofactors. (B) Following R-loop formation, Cas12a undergoes two Mg²⁺-dependent conformational changes, first to position the non-target strand and then the target strand in the RuvC active site. The two successive cleavage events are both carried out by the RuvC domain.

Successful PAM recognition and target binding results in DNA cleavage by Cas9 and Cas12a and formation of a double strand break (DSB) (Fig. 1) [2-4]. Cas9 generates DSBs in the PAM-proximal region using two metal-dependent nuclease domains, HNH and RuvC (Fig. 1A) [2,3]. The HNH domain employs a one-metal-ion mechanism to cleave the target strand, while the RuvC domain uses a two-metal-ion mechanism to cleave the non-target strand [21,22]. Cas12a uses a single RuvC nuclease domain to cleave both strands of the target DNA in the PAM-distal region (Fig. 1B) [23-25]. Both Cas9 and Cas12a must undergo large conformational changes to enable cleavage of both strands of DNA (Fig. 1). The Cas9 HNH domain only adopts a catalytic conformation upon formation of a full R-loop (Fig. 1A) [22,26-28]. Cas12a must rearrange the DNA substrate following non-target strand cleavage to allow access of the target strand within the RuvC active site (Fig. 1B) [23,25,29-32]. The conformational changes of both Cas9 and Cas12a are metal-ion dependent, underscoring the essential role of metal ions for Cas endonuclease function [27,30].

Recent studies have highlighted the impact of physiological metal ion conditions on Cas9 and Cas12a function. Some Cas9 orthologs are unable to bind DNA at the lower Mg²⁺ concentrations present in human cells [33]. Engineered Cas9 variants can overcome this limitation, allowing for improved genome editing efficiency. The RuvC domain of Cas9 is also sensitive to lower Mg²⁺ concentration that can be impacted by the presence of biomolecules that coordinate metal ions [34]. Inhibition of RuvC activity due to low Mg²⁺ availability results in the formation of a nick through cleavage by only the HNH domain. Cas12a is also prone to nicking at lower Mg²⁺ concentration [30], especially in the presence of target mismatches in the PAM-distal region [35-37]. At physiological Mg²⁺ concentrations, Cas12a cleavage of targets containing PAM-distal mismatches is significantly impaired [38]. Conversely, cleavage of targets containing seed mismatches or PAM mutations is relatively fast at lower Mg²⁺ concentrations [35].

Analyzing Cas9 and Cas12a kinetics under various metal ion conditions is essential for understanding their biochemical mechanisms and predicting off-target effects, leading to more accurate and safer gene editing technologies. In this chapter, we provide detailed protocols for performing in vitro cleavage assays that allow investigation of the impact of metal ions on Cas effector function, including both DNA binding and cleavage. The protocols provided in this chapter allow for rate measurements for the formation of both products containing cuts on one or both DNA strands using either negatively supercoiled or linear DNA substrates. Both sets of methods allow for detection of cleavage products without the need for expensive labeling techniques like radioactive ³²P or fluorescent dyes. Thus, these methods can be readily adapted for a wide variety of Cas effectors and targets.

2. Cleavage assays using negatively supercoiled plasmid DNA substrates

Negatively supercoiled plasmids are commonly used as substrates to measure DNA cleavage by Cas effectors. Cas9 and Cas12a generate double strand breaks, but may initially nick only one strand in the presence of target mutations or low metal ion conditions [30,34-39]. First-strand cleavage results in relaxation of negatively supercoiled substrates, while second-strand cleavage results in linearization of the plasmid (Fig. 2A). These distinct products have different mobilities during agarose gel electrophoresis, allowing for determination of the rates of both cleavage events. Here we describe standard protocols for cleavage assays using negatively supercoiled plasmid DNA as targets for Cas effector cleavage. We do not include methods for the in vitro synthesis of crRNA and trans activating CRISPR RNA (tracrRNA) or Cas9 purification, which have been described in detail in previous method papers [40,41]. We describe two distinct protocols for initiating DNA cleavage, either through the mixing of Cas9 RNA-protein complex (RNP) with the DNA substrate in the presence of Mg²⁺, or through the addition of Mg²⁺ to initiate cleavage following incubation of the Cas9 RNP and DNA. These two methods allow for differentiation between cleavage defects that may occur during or following DNA binding by the Cas effector, respectively. We also provide a third method for determining the rate of Cas effector binding to a DNA substrate in the absence of metal ions.

Figure 2: Negatively supercoiled target DNA substrates allow observation of first- and second-strand cleavage by Cas endonucleases.

(A) For a Cas9 target, cleavage of the target strand by the HNH domain results in relaxation of supercoils, which results in slower migration on an agarose gel. Cleavage of the non-target strand by the RuvC domain results in a double-strand break, converting the plasmid to a linear form that migrates between the relaxed and negatively supercoiled bands on the agarose gel.

(B-C) Design of oligonucleotides used for (B) a Cas9 target, derived from gene B of the λ phage genome or (C) a Cas12a target, derived from gene L of the λ phage genome. The oligonucleotides contain sticky ends when annealed together, allowing for ligation with doubly-digested plasmid to create the target plasmid.

2.1. Plasmid substrate preparation

Substrate plasmids can be generated using a restriction enzyme cloning protocol that we previously described [42]. Oligonucleotides containing a target sequence and PAM flanked by sticky ends for two restriction enzymes can be cloned into a plasmid digested with the corresponding restriction enzymes (Fig. 2B-C). For plasmid cleavage assays, we typically use pUC19 digested with BamHI and HindIII. After verifying the correct target plasmid sequence by Sanger or whole plasmid sequencing, larger quantities of the substrate plasmid are prepared. We typically use a Midi Kit and follow manufacturer’s directions outside of a few optimizations described below.

2.1.1. Equipment

Incubator at 37 °C

Shaker incubator at 37 °C

Spectrophotometer (e.g. NanoDrop from Thermo Fisher Scientific)

2.1.2. Buffers and Reagents

E. coli recA- competent cells (e.g. NEB 5-alpha from New England Biolabs)

HiSpeed Plasmid Midi Kit (QIAGEN)

2.1.3. Procedure

1. Transform 1 μL of the verified substrate plasmid into a recA- competent E. coli strain using standard protocols. Plate 100 μL of the recovery (or a dilution if necessary) on LB-agar plates containing the appropriate antibiotic. Incubate the plate overnight at 37 °C.

2. Using a single colony, inoculate 150 mL LB media supplemented with 100 μg/ml ampicillin. Grow the culture overnight at 37 °C with shaking (between 180-220 rpm).

3. Purify the plasmid DNA from the culture using a HiSpeed Plasmid Midi Kit (QIAGEN), following the manufacturer’s protocol with the following modifications:

   • Incubate the overnight culture on ice for at least 30 min prior to harvesting the cells. We find that this incubation ensures that all DNA remains negatively supercoiled upon lysis of the cells.
   • Elute the plasmid with 350 μl nuclease free water that was pre-incubated to 37 °C. This modification yields plasmid DNA with a higher concentration.

4. Determine the absorbance of the DNA at 260 nm using a spectrophotometer.

5. Dilute the plasmid stock to 100 ng/μL and aliquot plasmid into microcentrifuge tubes (50-100 μL each). Store at −20 °C. Each aliquot should be freeze-thawed no more than three times to ensure that plasmid remains negatively supercoiled.

2.2. Initiate cleavage by mixing Cas RNP and DNA in the presence of Mg²⁺

Cas9 must unwind and bind the target region of plasmid DNA prior to cleavage. The overall rate of unwinding, binding and cleavage can be measured in time course cleavage assays in which the DNA substrate and Cas effector-guide RNA complex are mixed together in the presence of metal ions to initiate cleavage (Fig. 3A) [21]. In these experiments, the Cas endonuclease protein is first incubated with crRNA (and trcrRNA if necessary) in a 1:1.5 molar ratio in cleavage buffer containing Mg²⁺ ion to form a ribonucleoprotein complex (RNP complex). Cas effectors are generally single turnover enzymes [12,16,43,44]. Therefore, the RNP complex concentration is used in excess over plasmid substrates to ensure complete cleavage. The reaction is initiated by adding negatively supercoiled plasmid into the reaction tube containing both RNP complex and Mg²⁺ ion. Aliquots of the reaction are removed at various time points and quenched by precipitating the Cas effector protein using phenol. The nucleic acid remains in the aqueous layer and can be extracted prior to analysis. The reaction is performed with different concentrations of Mg²⁺ added to the reaction buffer to determine the effect of divalent metal ions on the overall rate of cleavage [35].

Figure 3: Cleavage of negatively supercoiled DNA substrate when initiating by mixing DNA and Cas RNP in the presence of metal ions.

(A) Schematic of the cleavage assay. The Cas effector and RNA are first incubated in the presence of Mg²⁺ to prepare the Cas RNP. The DNA substrate is also diluted in a buffer containing Mg²⁺. Both components are mixed to initiate DNA binding and cleavage. The rates determined for the cleavage of each strand encompasses all events that must occur prior to cleavage, including PAM searching, DNA unwinding and R-loop formation, and conformational rearrangements.

(B) Agarose gel showing a Cas9 cleavage time course for the gene B target at 10 and 1 mM MgCl₂. Nicked DNA (ni), linear DNA (li), and negatively supercoiled DNA (nSC) bands are labeled. The time points were 7 s, 15 s, 30 s, 1 min, 2 min, 5 min, 15 min, and 30 min. The two controls are DNA alone (−) and DNA with Cas9 without crRNA and tracrRNA (−cr). Both controls were performed at the higher MgCl₂ concentration for the longest time point. The gel was post-stained with SybrSafe for 1 h before imaging.

(C-D) Images showing steps of quantification of the agarose gel in (B) using ImageJ. (C) Following background subtraction, lanes were selected using the rectangle tool. (D) The bands were plotted and integrated using the magic wand tool. The area under the curve reported in the Results window were used to determine the fraction of DNA that had undergone first or second-strand cleavage.

(E-F) Rate curves plotting fraction cleaved versus time for (E) first- and (F) second-strand cleavage at the two MgCl₂ concentrations. The data in (E) were fit to a double-exponential equation, while the data in (F) fit better to a single-exponential. Curve fitting was performed in GraphPad Prism. The average of three replicates is shown and error bars represent standard deviation. Data are adapted from [34].

To analyze the rate of cleavage, DNA products are separated by agarose gel electrophoresis (Fig. 3B). The gels are stained with SYBR Safe following electrophoresis to allow visualization of negatively supercoiled, relaxed, and linear DNA. Post-staining the gel is essential to ensure proper migration of the three DNA species, whose migration can be affected when DNA intercalators are present in the gel matrix. Following gel visualization, the intensity of each DNA band can be quantified using ImageJ software (Fig. 3C-D) [45,46]. The relaxed DNA band contains products that have been nicked on a single strand through a first-strand cleavage event, while the linear DNA band contains products that underwent second-strand cleavage. Thus, separation of a relaxed and linear product enables determination of the rate of both a first- and second-strand cleavage event (Fig. 3E-F) [35].

Cas9 has previously been observed to display biphasic cleavage kinetics [21,47]. The two phases of cleavage have recently been linked to conformational differences that occur upon DNA binding that result in different cleavage timescales for a Cas9d ortholog [48]. If such biphasic kinetics are observed, it is appropriate to fit the kinetic data to a double-exponential rate equation that estimates the rate constants for the fast and the slow cleavage phase [47]. However, we sometimes observe that the slow phase cannot be accurately determined by the graphing software. In these cases, it is appropriate to fit the data to a single-exponential rate equation that models a single cleavage phase. In either case, the rate constants represent the combined rates of all events that lead to cleavage, including PAM searching, target unwinding and accompanying conformational changes, and cleavage catalysis.

We have included exemplar data for a Cas9 target derived from the gene B region of the λ phage genome (Fig. 2B) [34]. While first-strand cleavage rates were observed to be biphasic, and therefore fit to a double-exponential rate equation (Fig. 3E), second-strand cleavage rates were observed to have a single phase, and were therefore fit to a single-exponential rate equation (Fig. 3F). We observed a similar rate of first-strand cleavage between the two Mg²⁺ conditions tested (Fig. 3E), with some differences between the distribution and rates of the fast and slow phase. In contrast, we observed much slower second-strand cleavage at 1 mM Mg²⁺ in comparison to 10 mM Mg²⁺ (Fig. 3F), suggesting that cleavage by one of the two Cas9 nuclease domains is impaired at lower metal ion conditions. In our previous study, we observed that the slow second-strand cleavage kinetics are due to slower cleavage by the RuvC domain at lower metal ion conditions [34].

In this section, we provide detailed methods for performing the cleavage assay initiated by mixing DNA and Cas RNP together in the presence of MgCl₂, tips for how best to separate and visualize the products via agarose gel electrophoresis, and a description of how we perform analysis of the data using ImageJ and GraphPad Prism.

2.2.1. Equipment

• Heat block for 37 °C incubation

• Microcentrifuge

• Agarose gel apparatus (e.g. Corning Axygen 15 cm Horizontal Gel Box)

• Agarose gel casting kit with 15 cm gel tray and 30-well 1.5 mm comb

• Power supply

• Orbital shaker

• Agarose gel imaging system (e.g. Bio-Rad Gel Doc EZ)

• Staining tray

2.2.1. Buffers and reagents

• 5X reaction buffer (100 mM HEPES, pH 7.5, 500 mM KCl, 5 mM DTT, 50 mM or 5 mM MgCl₂ and 25% glycerol)

• 25:24:1 phenol-chloroform-isoamyl alcohol (Invitrogen)

• 2X DNA loading buffer (50 mM EDTA, 0.1% bromophenol blue, 0.1% xylene cyanol FF, 10% glycerol).

• TAE buffer (40 mM Tris base, 20 mM acetic acid, and 1 mM EDTA)

• Agarose

• SYBR Safe (Thermo Fisher Scientific)

2.2.2. Procedure

2.2.2.1. Perform Cas9 plasmid DNA cleavage time course

1. We typically perform eight time points at 7 s, 15 s, 30 s, 1 min, 2 min, 5 min, 15 min, and 30 min, although these may vary depending on the rate of cleavage for a given target. For each time point, we prepare two tubes, one containing 10 μL of phenol-chloroform-isoamyl alcohol (for time point quenching) and one containing 10 μL of 2X DNA dye (for DNA extraction). Arrange each set of tubes in order of time points to ensure tubes are ready for quenching time points.

2. We typically perform cleavage reactions at two different MgCl₂ concentrations, 10 and 1 mM. Reactions can be set up in parallel by adjusting the concentration of MgCl₂ in the reaction buffer. Initiation of the reactions should be staggered to ensure that all time points can be taken.

3. Cas9, crRNA and tracrRNA can be prepared as previously described [refs]. On ice, dilute Cas9 to 1 μM with 1X reaction buffer chilled on ice. On ice, dilute in vitro transcribed crRNA and tracrRNA with 5X reaction buffer and nuclease free water to a final concentration of 1 μM RNA and 1X reaction buffer. Leave the diluted reagents on ice.

4. To form RNP complex, mix 100 nM Cas9 with 150 nM crRNA and 150 nM tracrRNA in pre-chilled 1X reaction buffer 1 or reaction buffer 2 in a final volume of 50 μL. Leave the RNP on ice until all reagents are prepared.

5. Dilute the plasmid target with pre-chilled 5X reaction buffer and nuclease-free water to a final volume of 50 μL and a final concentration of 15 ng/mL plasmid and 1X reaction buffer. Leave the DNA on ice until all reagents are prepared.

6. Incubate the diluted RNP complex and plasmid DNA at 37 °C for 10 min.

7. Add the contents of the DNA tube to the RNP tube (or vice versa) to initiate the cleavage reaction. Mix well by pipetting up and down 3-4 times. Continue to incubate the reaction at 37 °C.

8. Remove 10 μl of sample at each time-point and rapidly mix with the aliquoted phenol-chloroform-isoamyl alcohol in the appropriate time point tube to quench the reaction. Pipet up and down 5-6 times to ensure mixing.

   • Tip: It is necessary to have a pipet ready with a tip on it to take a time point immediately after initiating cleavage. We typically find that 7 s is the shortest amount of time in which the reaction components can be adequately mixed, and a time point quenched when time points are performed manually.

9. Additional tubes should be prepared as controls. We generally perform one control in which DNA is incubated in reaction buffer without RNP and another in which DNA and Cas9 are incubated without the RNAs. Both controls would be incubated at 37 °C for the longest time point.

2.2.2.2. Prepare samples for running agarose gel

1. Prepare a 1% agarose gel in TAE buffer using standard protocols. Note that it is important to not add any dye to the gel to ensure the migration of DNA on the gel based on the topology of the DNA. We typically use a 15 cm gel tray and one or two 30-well combs (1.5 mm thickness), depending on how many samples will be run. If using two combs, one is placed at the top of the gel and one in the middle. Different sets of samples can be run in each set of wells. Note that the gel can be prepared prior to performing the time course assay.

2. Centrifuge time point samples in a microcentrifuge at the highest speed for 1 min and then remove 9 μl of DNA from the top aqueous layer into the prepared 2X DNA dye tubes.

   • Tip: Removal of the aqueous layer can be facilitated by adding a small amount (2-5 μL) of 2X DNA dye to the quenched samples prior to centrifugation. The dye will separate into the organic layer and make the aqueous layer more visually distinct. Note that addition of the DNA dye will dilute the aqueous layer.

3. Load 15 μL of each sample into wells and run gel at 110 V for 1 hour 30 min to ensure good separation between nicked and linearized products. Run a DNA ladder (e.g. Generuler 1kb DNA ladder from Thermo Fisher Scientific) in at least one lane adjacent to the time point samples.

4. Dilute 20 μL of SYBR safe into 200 mL of TAE buffer. Stain the gel with SYBR safe for 1 hour while shaking gently on an orbital shaker.

5. Image the gel using a gel imager equipped with a UV transilluminator. Export the gel image in a format that is compatible with ImageJ (e.g. TIFF).

2.2.2.3. Quantification of DNA cleavage versus time

1. Gels should have DNA bands corresponding to negatively supercoiled and linear DNA, and may also have relaxed DNA if DNA nicking occurred (Fig. 3B). Controls (e.g. nicked and linearized plasmids prepared using restriction enzymes) can be run to ensure that the migration of DNA bands corresponds to nicked and linearized products.

2. Determine the intensity of each DNA band using Image J (Fig. 3C-D).

   • Open the exported gel image in ImageJ.
   • Ensure that the gel image is horizontal. Adjust using image rotation tools if necessary.
   • Invert the image using Edit > Invert to produce dark DNA bands on a light background.
   • Perform background subtraction using the Process > Subtract Background… dialog. We use the “Light background” setting with a rolling ball radius of 25.
   • Using the Rectangle tool from the tool bar, draw a rectangle around the first lane containing a time point sample (Fig. 3C). Use the keyboard shortcut control-1 on Windows or command-1 on Mac to select the first lane.
   • Drag the rectangle. This should create a new rectangle that can be positioned on the next lane. Use the keyboard shortcut control-2 on Windows or command-2 on Mac to select the next lane. Repeat this for the remaining lanes.
   • Once all lanes have been outlined with rectangles, use the keyboard shortcut control-3 on Windows or command-3 on Mac to plot the lanes. This will bring up a new window containing 2D plots of DNA band peaks.
   • Use the magic wand tool to determine the area under each peak (Fig. 3D). Lines can be drawn using the line tool from the tool bar to ensure an even baseline and to distinguish between different peaks.
   • Copy the peak intensity data to a worksheet in Excel, keeping track of values from the relaxed, linear, and supercoiled peaks (Fig. 3D, inset on right).

3. In Excel, calculate the intensity of total DNA in each sample by adding the intensities of the three bands.

4. First-strand cleavage describes the rate at which the DNA is nicked based on an initial cleavage event. Nicked DNA is eventually linearized, so the fraction of DNA that underwent the first cleavage event can be determined by adding the intensity of the nicked and linear products.

   $$fractionofDNAthatunderwentfirststrandcleavage=\frac{(Inicked+Ilinear)}{Iall}$$

   where I nicked is intensity of relaxed DNA band, I linear is intensity of linear DNA band, and I all is intensity of all bands

5. DNA that has undergone second strand cleavage should be linearized, and can be determined using:

   $$fractionofDNAthatunderwentsecondstrandcleavage=\frac{Ilinear}{Iall}$$

   where I linear is intensity of linear band and I all is intensity of all bands.

6. Rate constants can be derived from fits of individual replicates to either a single-exponential or a double-exponential rate equation in a graphing software. We typically use GraphPad Prism, using the built-in “One-phase association” non-linear regression for single-exponential rates or the “Two-phase association” for double-exponential rates.

   • The equation for One-phase association is:

     $$y=y_0+(plateau-y_0)\times (1-e^{(-kx)})$$
   • The equation for Two-phase association is:

     $$y=y_0+SpanFast\ast (1-e^{(-k_{fast}x)})+SpanSlow\ast (1-e^{(-k_{slow}x)})$$
     where $y_0$ is the $\mathrm{y}$ value at time 0, $plateau$ is the amplitude of the curve, and $k$ is the observed rate constant. We constrain $y_0$ to 0 and $plateau$ to less than 1. SpanFast and SpanSlow estimate the fraction of the signal that is due to the fast or slow phase, respectively.

7. All experiments should be performed at least three times. If rate constants are to be reported, fit each replicate individually to the appropriate rate equation and determine the average and standard deviation of the rate constants determined for each replicate. We depict the rate curves using the average fraction cleaved at each time points, with error bars representing the standard deviation for fraction cleaved at each time point (Fig. 3E-F).

2.3. Initiate cleavage through the addition of Mg²⁺ ion to pre-bound RNP-DNA complex

As described above, cleavage reactions that are initiated by mixing the RNP and DNA in the presence of Mg²⁺ report on the rate of all events leading up to, and including, DNA cleavage. As an alternative, incubation of the RNP and plasmid DNA prior to the addition of Mg²⁺ allows for formation of an R-loop prior to initiation of cleavage (Fig. 4A). Performing the cleavage assay in this manner eliminates the rate of PAM searching, DNA unwinding and target binding from the overall rate of the cleavage reaction (Fig. 1) [35,47]. Thus, cleavage rates and rate constants reflect the rate of metal-dependent steps that occur after DNA binding, including conformational changes of endonuclease domains and cleavage catalysis.

Figure 4: Cleavage of negatively supercoiled DNA substrate when initiating by adding metal ions.

(A) Schematic of the cleavage assay. The Cas RNP and DNA substrate are first incubated in the absence of Mg²⁺. Cleavage is initiated by adding Mg²⁺. The rates determined for the cleavage of each strand encompasses Mg²⁺-dependent events that occur following R-loop formation, including conformational rearrangements and cleavage catalysis.

(B) Agarose gel showing a Cas9 cleavage time course for the gene B target initiated with either 10 and 1 mM MgCl₂. The time points and labels are as in Fig. 3B.

(C-D) Rate curves plotting fraction cleaved versus time for two different initiation methods at (C) 10 mM MgCl₂ or (D) 1 mM MgCl₂. First-strand cleavage data were fit to double-exponential equations while second-strand cleavage data were fit to single-exponential equations in GraphPad Prism. The average of three replicates is shown and error bars represent standard deviation.

In Figure 4, we have included example data for the λ phage gene B Cas9 target comparing rates when cleavage was initiated with Mg²⁺ with the data shown in Figure 3E-F. We observed a large increase in rate of first-strand cleavage at both MgCl₂ concentrations when cleavage was initiated by adding Mg²⁺ in comparison to when cleavage was initiated by adding DNA (Fig. 4C-D). This difference reflects the slow rate of DNA binding, which limits the rate of cleavage when the reaction is initiated by adding DNA. The difference in rates of first-strand cleavage was larger at 10 mM MgCl₂ (Fig. 4C) in comparison to 1 mM MgCl₂ (Fig. 4D), consistent with previous observations of slower conformational kinetics of the HNH domain at lower divalent metal ion conditions [27]. Second-strand cleavage rates were similar regardless of initiation methods (Fig. 4C-D), suggesting that pre-binding the Cas9 RNP to DNA has little effect on the rate of cleavage by the RuvC domain [34].

In this section, we describe a protocol for the measurement of plasmid DNA cleavage rate by initiating the reaction with Mg²⁺ ions (Fig. 4). All equipment and reagents are the same as for the procedure described in section 2.2. with the noted exceptions.

2.3.1. Procedure

1. Prepare a 5X reaction buffer without Mg²⁺ containing 100 mM HEPES, pH 7.5, 500 mM KCl, 5 mM DTT, and 25% glycerol.

2. Prepare 40 μL of 5 mM or 50 mM MgCl₂ solution in 1X reaction buffer.

3. Prepare tubes for time points containing phenol solution and 2X DNA dye as described in section 2.2.2.1.

4. Prepare the RNP and DNA as described in section 2.2.2.1 using the reaction buffer lacking Mg²⁺.

5. Incubate the diluted RNP, plasmid and MgCl₂ solutions for 10 min at 37 °C.

6. Mix 40 μL of the RNP and the DNA to allow for DNA binding. Mix well by pipetting up and down 3-4 times and incubate for 30 min at 37 °C.

7. Add 20 μL of the MgCl₂ solution into the reaction tube and mix well by pipetting up and down 3-4 times.

8. Quench time points, prepare samples for gels, and analyze results as described in sections 2.2.2.1-2.2.2.3.

2.4. DNA binding time-course analyzed by determining fraction cleaved

We have previously observed that Mg²⁺ may decrease the rate of Cas effector binding to negatively supercoiled plasmids [35]. These observations prompted us to develop an assay that enables the measurement of the fraction of DNA that is bound at various time points in the absence of Mg²⁺. Our assay makes use of the observation that for most DNA targets, Cas effectors cleave the DNA rapidly following target binding in the presence of high Mg²⁺ concentrations [21,35]. Thus, addition of Mg²⁺ for a brief period allows for cleavage of all of the DNA that has been bound by RNP. Measurement of the fraction cleaved provides information on the fraction of DNA that was bound by RNP at a given time point.

To optimize this assay, we first determined how long it was necessary to mix an RNP-DNA complex with Mg²⁺ to see complete cleavage. We found that a Cas12a RNP-DNA complex formed using a reaction buffer that lacked Mg²⁺ for 30 min at 37 °C, could fully cleave the DNA upon mixing with 10 mM MgCl₂ for 5 s. This result indicates that any RNP that is bound to DNA would be able to fully cleave within 5 s upon addition of Mg²⁺. Importantly, the amount of time needed to fully cleave the DNA upon addition of Mg²⁺ may vary between targets, so it is important to perform this control for every DNA target tested.

Using this observation, we developed a protocol for monitoring the fraction of Cas RNP that is bound to DNA in the absence of Mg²⁺ over a time course (Fig. 5A) [35]. In this assay, the RNP and DNA are mixed in the absence of Mg²⁺ and incubated at 37 °C. At a given time point, an aliquot is removed from this binding reaction and mixed with Mg²⁺ at a final concentration of 10 mM. Cleavage is allowed to proceed for 5 seconds, and then the RNP-DNA is transferred to a phenol solution for quenching. The fraction cleaved is determined and used to estimate the fraction of DNA that was bound by the RNP at a given time point.

Figure 5: Measuring the rate of DNA binding in the absence of metal ions.

(A) Schematic of the binding assay. Binding is initiated by mixing the Cas RNP and DNA substrate in the absence of Mg²⁺. At various time points, aliquots are removed from the binding reaction and mixed for 5 s with 10 mM MgCl₂ and then immediately quenched. All DNA that is bound by the Cas RNP at that time point will be cleaved, while DNA that remains unbound will remain negatively supercoiled.

(B) Agarose gel showing a Cas12a cleavage time course for the gene L A2T target. The first two cleavage reactions were performed by initiating with DNA in the presence of 10 or 1 mM MgCl₂. The third reaction is the binding reaction in the absence of MgCl₂. The time points and labels are as in Fig. 3B.

(C) Rate curves plotting fraction cleaved versus time for the three different MgCl₂ conditions that were present during DNA binding. The first-strand cleavage rate is plotted, as binding is rate limiting for first-strand cleavage for the gene L A2T target [35]. The top graph shows all time points, while the bottom graph shows only the early time points. The average of three replicates is shown and error bars represent standard deviation. Data are adapted from [35].

We previously used this technique to measure the rate of cleavage of a Cas12a DNA target derived from gene L of the λ phage genome (Fig. 2C, 5B-C) [35]. This target contains an A2T mutation within the seed region. The seed mismatch slows the rate of Cas12a DNA binding, especially at higher Mg²⁺ concentrations, resulting in DNA binding being rate-limiting. Thus, measurement of the rate of cleavage using the procedures described in section 2.2 allowed us to estimate the rate of binding at variable Mg²⁺ concentrations. By using the procedure described below, we were also able to estimate a rate of Cas12a binding to the gene L A2T target in the absence of metal ions. We observed the fastest binding of Cas12a to the target when metal ions were absent during binding and the slowest rate of binding at the highest Mg²⁺ concentration tested (Fig. 5B-C).

Below we describe the in-depth procedure for measuring the rate of target binding in the absence of metal ions (Fig. 5). All equipment and reagents are the same as for the procedure described in section 2.2. with the noted exceptions.

2.4.1. Procedure

1. Prepare a 5X reaction buffer without Mg²⁺ containing 100 mM HEPES, pH 7.5, 500 mM KCl, 5 mM DTT, and 25% glycerol.

2. Prepare a 50 mM MgCl₂ solution in 1X reaction buffer. Prepare tubes containing 2.5 μL of this solution, one for each time point to be tested.

3. Prepare additional tubes for time points containing phenol solution and 2X DNA dye as described in section 2.2.2.1.

4. Prepare the RNP and DNA as described in section 2.2.2.1 using the reaction buffer lacking Mg²⁺.

5. Incubate the diluted RNP and plasmid solutions for 10 min at 37 °C.

6. Mix the contents of the RNP tube with the contents of the DNA tube to allow for DNA binding. Mix well by pipetting up and down 3-4 times. Continue incubating at 37 °C.

7. At each time point, remove 10 μL of the RNP-DNA mixture and mix rapidly by pipetting up and down for 5 s (or the appropriate time determined for individual DNA targets) to ensure complete cleavage of all bound DNA. At the end of the 5 s, using the same pipet tip, immediately move 10 μL of the RNP-DNA-Mg²⁺ mixture to the phenol-containing tube to quench both DNA binding and cleavage. Repeat this process for the remaining time points.

8. Prepare samples for gels, and analyze results as described in sections 2.2.2.1-2.2.2.3.

3. Cleavage assays using linear DNA substrates

While negatively supercoiled DNA provides a convenient means to monitor nicking events on both the strands of the DNA target, the topology of supercoiled DNA can have significant effects on the ability of Cas effectors to unwind and bind DNA. Linear DNA targets are relevant to the native biology of CRISPR-Cas effectors, as the genomic DNA of bacteriophages is linear at various stages of the viral life cycle [49]. Thus, it is also useful to be able to compare cleavage between supercoiled and linear DNA targets.

Short DNA oligonucleotides are commonly used as linear substrates for Cas effector cleavage [2,47]. For these substrates, individual strands of a double-stranded target can be labeled differentially, either by the addition of different fluorophores to each strand or by labeling each strand radioactively with ³²P and performing separate cleavage assays to monitor cleavage on each strand. Although very useful, these labeling techniques can become cost prohibitive, especially when testing a large number of different target sequences and with the rising costs of radioactive ATP. Additionally, short DNA oligonucleotides do not correspond well to the longer linear DNA that Cas effectors would encounter during their native function.

Linearized plasmids or PCR products containing target sequences can serve as alternative substrates that better represent target sequences within the context of longer linear DNA [4]. However, when using a longer linear DNA substrate, it is difficult to observe all cleavage products by gel electrophoresis. Although products that have undergone cleavage events on both strands are readily separated due to the smaller size of the resulting fragments, uncleaved linear DNA and nicked fragments have similar migration patterns and are difficult to separate. This prevents quantification of first-strand DNA cleavage when using linear DNA in the absence of labels on each DNA strand.

To observe all the products from the first and second strand cleavage events in a cleavage assay with linear DNA substrate, we adapted a method using denaturing polyacrylamide gel electrophoresis [50]. This method relies on the formation of a nick toward the center of a linear DNA fragment (Fig. 6A). When the linear DNA undergoes a first-strand cleavage event, a hinge is formed at the site of nick. Urea disrupts base stacking interactions between base pairs on either side of the nick, allowing bending of the nicked DNA and slower migration on the gel. DNA can be visualized using SYBR Gold stain to avoid the need for labels within the DNA prior to cleavage.

Figure 6: Cleavage of long linear target DNA substrates.

(A) Schematic showing strategy for separation of linear target DNA substrates following first- and second-strand cleavage events. A 600-700 bp fragment can be PCR amplified from a target plasmid. The target should be at the center of the PCR product. First-strand cleavage allows for DNA bending under denaturing conditions, which results in slower migration on a polyacrylamide gel. Second-strand cleavage results in two DNA products of the same length, which co-migrate on the gel and can be visualized readily by SYBR Gold stain.

(B) Cleavage of a ~650 bp PCR product using the Cas9 D10A variant, in which only the HNH domain is able to cleave the target, producing a nicked product. A denaturing Urea-PAGE gel enabled separation of the nicked and uncleaved DNA.

(C) Cleavage of a ~650 bp PCR product using Cas12a. The target contained a PAM-distal mutation, which results in slow second-strand cleavage and accumulation of a nicked product. A similar length PCR product cleaved by D10A and WT Cas9 were included as controls to ensure that the slowly migrating species is consistent with the migration of a nicked DNA. Gels were visualized using SYBR Gold staining.

3.1. Linear substrate preparation

To allow for a direct comparison of cleavage between negatively supercoiled and linear substrates, we recommend PCR amplifying Cas9 or Cas12a targets using the target plasmids described in section 2.1.3 as templates (Fig. 6A). In the development of this method, we PCR amplified DNA targets of various lengths and with targets located in the center of the DNA or toward one of the ends. We found that having the cleavage site at the center of a 600-700 bp fragment provided the best separation between a nicked DNA product and the uncleaved linear DNA. In addition, placing the target at the center of the PCR product results in two shorter fragments of the same length upon second strand cleavage, which improves the visualization of the shorter fragments when staining with SYBR Gold. The procedure below outlines a general framework for the preparation of these substrates.

3.1.1. Equipment

• Thermal cycler

• Agarose gel apparatus

• Spectrophotemeter (e.g. Thermo Fisher NanoDrop)

3.1.2. Buffers and reagents

• DNA polymerase for PCR (e.g. New England Biolabs Q5 High-Fidelity DNA polymerase)

• Target plasmid (prepared as described in section 2.1.3)

• PCR primers (designed as described below)

• dNTPs

• Agarose gel with dye for detecting DNA

• PCR cleanup kit (e.g. Promega Wizard Kit or Qiagen QIAquick PCR Purification kit)

• Nuclease-free water

3.3. Procedure

3.3.1. Generation of PCR amplified DNA substrate

1. Prepare a DNA substrate in the range of 600-700 bp by PCR amplifying the target pUC19 plasmids prepared as described in section 2.1.3. Primers for PCR should be designed in such a way that the target site of the Cas proteins is in the middle of the substrate (Fig. 6A). We typically perform a 200 μL reaction using 30 cycles of a standard PCR protocol.

2. Confirm the length of the PCR product by running a small amount on an agarose gel.

3. Purify the PCR products using a standard PCR cleanup kit following the manufacturer’s instructions. Elute the DNA using 30 μL of nuclease-free water.

4. Determine the concentration of the DNA using a spectrophotometer. Dilute the DNA to 100 nM final concentration using nuclease-free water.

5. Aliquot the DNA and store at −20 °C.

3.2. Cleavage assays and separation of nicked products for linear substrates

To verify that we could separate nicked products from uncleaved linear DNA, we used a Cas9 HNH nickase in which the RuvC active site is mutated (D10A) such that only the HNH domain is active [2]. Cleavage by the HNH domain results in nicking of only the target strand of the DNA and ensures a nicked product. We performed cleavage of a Cas9 target using the HNH nickase and were able to separate nicked DNA products from the uncleaved DNA by denaturing polyacrylamide gel electrophoresis (Fig. 6B). Having confirmed that nicked products can be separated through this method, we next tested cleavage of a Cas12a target containing a PAM-distal mismatch. We have previously observed that PAM-distal mismatches cause slower second-strand cleavage by Cas12a, resulting in a nicked product when using negatively supercoiled DNA substrates [35,36,38]. Similarly, using a linear PCR product as substrate, we observed a DNA product with a similar migration to the Cas9 HNH nickase product (Fig. 6C), indicating that Cas12a also nicks linear DNA containing PAM-distal mismatches.

The cleavage reactions for linear substrates can be performed similarly to procedures described in sections 2.2.2.1, 2.3.1 or 2.4.1, with a few modifications noted below. The major differences in the below protocols are in the methods by which the DNA is separated and analyzed.

3.2.1. Equipment

• Polyacrylamide gel electrophoresis equipment

• High voltage power supply (e.g. Bio-Rad PowerPac HC)

• Gel imager (e.g. Cytiva Typhoon imager)

3.2.2. Buffers and reagents

• 10X TBE buffer (pH 8.3): 1M Tris Base, 1M Boric acid, 0.02M EDTA

• SYBR Gold Nucleic Acid Gel Stain (Thermo Fisher Scientific)

3.2.3. Procedure

1. For cleavage assays, we typically use a final DNA concentration of 5 nM in the reaction, resulting in a final DNA concentration of 2.5 nM following quenching and dilution with 2X DNA dye. This concentration is sufficient to allow for visualization of the DNA using SYBR Gold stain.

2. With the exception of DNA concentration, cleavage reactions can be performed as described in sections above. For the accompanying data (Fig. 6B-C), we initiated cleavage by mixing RNP and DNA together in the presence of MgCl₂ as described in section 2.2.2.1. Time points, quenching, DNA extraction, and preparation of samples for gel electrophoresis can be performed as described in section 2.2.2.1.

3. Prepare an 8% denaturing PAGE gel in 1X TBE with 7M urea. To ensure visualization of all DNA fragments on the gel, we recommend using an apparatus with gel lengths of 28 cm or longer and spacers of 0.75 mm or thinner.

4. Load 10uL from each reaction to the wells of the TBE PAGE gel.

   • Tip: Do not heat the sample before loading on the gel. The DNA should remain double-stranded while running on the gel, but the denaturant will serve to disrupt stacking interactions at the nick site and alter the migration of nicked DNA.
5. Separate the cleavage reaction products at 18W constant current at ambient temperature.

   • Tip: Optimal separation of the nicked and uncleaved DNA requires higher resolution afforded by longer and thinner gels. We run our 28 cm gels for 1 hour and 45 min until the bromophenol blue dye front has migrated ~3/4 of the gel length.

6. Dilute the SYBR Gold (10,000X) by adding 2 μL of the dye to 100 mL of distilled water. Remove the gel from the apparatus and plates and carefully transfer to a staining tray containing the diluted SYBR Gold for 10 minutes.

7. Image the gel using a gel imager. Gels can be quantified as described above in section 2.2.2.3.

4. Summary and conclusion

In this chapter, we described methods for the analysis of the effect of divalent metal ions on the DNA binding and cleavage activities of Cas effectors. The methods described above allow for detection of the two separate cleavage events by the HNH and RuvC domains of Cas9, or the two successive cleavage events by the single RuvC domain of Cas12 proteins. We demonstrated that nicked fragments can be successfully separated and quantified when using either negatively supercoiled or linear DNA substrates. These methods allow for the determination of rates of the first cleavage event by various Cas effectors. First-strand cleavage rates are especially important when using lower Mg²⁺ concentrations, which have been shown to cause defects in RuvC cleavage by Cas9 and in second-strand cleavage by Cas12a. Initiating cleavage through three separate methods allows for differentiation between binding and cleavage defects, and enables the monitoring of the rate of DNA binding in the absence or presence of Mg²⁺. Overall, these methods allow for detailed examination of the impact of divalent metal ions on distinct steps of Cas effector activity.