Crystal structure of the Cas1–Cas2 complex

To gain insights into the structural organization of the Cas1–Cas2 complex, we determined the crystal structure of the complex. Crystal structures of Cas1 and Cas2 alone from various organisms, including E. coli K12, have been reported^{22–26,29,30}. Cas1 proteins are asymmetrical homodimers with each monomer having an N-terminal β-sheet domain and C-terminal α-helical domain^23,24,26. Cas2 proteins are symmetrical homodimers with a core ferredoxin fold^22,25,29,30. We purified each protein and reconstituted the complex in vitro. Gel filtration chromatography showed the co-migration of both proteins as one peak, confirmed by SDS-PAGE analysis of the peak fractions (Supplementary Fig. 1d,e). The Cas1–Cas2 complex yielded crystals that diffracted X-rays to 2.3-Å resolution. We determined the structure by single-wavelength anomalous dispersion (SAD) using selenomethionine-derivatized crystals and refined the resulting model to an R_work/R_free of 22%/24% (Table 1).

Table 1

Data collection and refinement statistics.

	Native	Se derivative
Data collection
Space group	P21	P21
Cell dimensions
a, b, c (Å)	94.875, 125.70, 99.31	93.70, 127.70, 99.32
a, b, g (°)	90, 102.74, 90	90, 102.32, 90
Resolution (Å)	62.85–2.3 (2.383 – 2.3)	48.63 – 2.89
Rsym or Rmerge	0.1301 (1.088)	0.2045 (1.576)
I / sI	9.19 (1.00)	10.32 (1.37)
Completeness (%)	97.22 (91.21)	91.87
Redundancy	3.1 (2.9)	7.8 (7.2)
Refinement
Resolution (Å)	62.85 – 2.3 (2.383 – 2.3)
No. reflections	97,929 (9,271)
Rwork / Rfree	0.225/0.245
No. atoms	10,451
Protein	9,926
Water	525
B factors
Protein	55.60
Water	53.90
r.m.s. deviations
Bond lengths (Å)	0.003
Bond angles (°)	0.62

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^*One crystal was used per data set. Values in parentheses denote highest-resolution shell.

The overall architecture of the asymmetric unit is a heterohexameric complex consisting of two Cas1 dimers (Cas1a-b and Cas1c-d) that sandwich one Cas2 dimer (Fig. 2a). Cas1a and Cas1c make contacts with the Cas2 dimer and no contacts are observed between Cas1b or Cas1d and the Cas2 dimer. The Cas1c–Cas2 protein-protein interface buries a large surface area of ~3,100 Ų, whereas the Cas1a–Cas2 interface buries an additional 800 Ų contributed by the C-terminus of Cas1a, as described further below. Superposition of the two Cas1 dimers (a-b dimer with c-d dimer) shows high structural similarity, with a root mean square deviation (r.m.s.d.) of 0.394 Å for the C-alpha atoms (Supplementary Fig. 2a,b). Similar contacts are present between Cas1a and Cas1c with Cas2 on opposite sides, creating a symmetrical complex. While Cas1 and Cas2 predominantly form a heterotetrameric complex in solution, our crystal structure suggests the complex may also be capable of accessing a hexameric state during acquisition.

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Figure 2

Crystal structure of the Cas1–Cas2 complex

(a) The overall structure consists of a Cas2 dimer (yellow and orange) and two Cas1 dimers (a-d, blue and teal). (b) Superposition of the Cas1a-b dimer with the previously determined E. coli Cas1 structure (gray, PDB 3NKD²⁴). The dashed orange circle highlights the conformational change observed in the a-helical domain of Cas1a. (c) Superposition of the Cas2 dimer in the complex with the previously determined E. coli Cas2 structure (gray, PDB 4MAK³⁰). The arrows point to the last resolved residue in the 4MAK structure. The N and C indicate the termini of each monomer; root-mean-square deviations (r.m.s.d.) of the superpositions are indicated.

Conformational changes and contacts within the complex

The interface between Cas1 and Cas2 consists of hydrogen bonding, electrostatic and hydrophobic interactions. We observe extensive electrostatic contacts between three arginine residues (R245, R252, R256) in α8 of Cas1 with two acidic residues (E65 and D84) of Cas2 (Fig. 3a and Supplementary Fig. 2c). The R252 residue is positioned between E65 and D84 and may sample salt bridges between the two acidic residues, although we observe continuous density between R252 and E65 at the Cas1a–Cas2 interface. In the same region, backbone hydrogen-bond contacts are present between the newly resolved Cas2 β7 C-terminus and β4 of Cas1 as discussed further below.

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Figure 3

Disruption of complex formation affects spacer acquisition in vivo

(a) A close-up view of the Cas1a–Cas2 protein-protein interface, with annotations for the residues involved in electrostatic interactions. (b) View of the ordered C-terminal tail of Cas1a with the electron density mesh contoured at 1.0 sigma. (c–e) Agarose gels of in vivo acquisition assays with mutations of Cas1 and Cas2 at the C-termini (c) and the electrostatic interface (d,e). (f) Western blot of FLAG immunoprecipitations in BL21-AI cells expressing Cas1-FLAG and Cas2-HA or various mutations of Cas1 and Cas2. Despite the low expression of Cas2 E65R, we still detect its co-elution with Cas1.

To identify Cas1 and Cas2 conformational changes that occur upon complex formation, we superimposed previously determined structures of apo Cas1 (PDB 3NKD)²⁴ and Cas2 (PDB 4MAK)³⁰ from E. coli with the Cas1–Cas2 complex structure (Fig. 2b, c). In addition to minor conformational changes present in the canonical βαββαβ ferredoxin fold of Cas2, the C-terminus forms two antiparallel β-sheets (β6–β7) that contact β4 of Cas1 (Fig. 2c and Fig. 3a). This region is unresolved in the apo-Cas2 structure, which terminates at the C-terminus of β5. Presumably, the β6–β7 region is flexible prior to complex formation with Cas1.

Although Cas1 does not undergo major conformational changes upon Cas2 binding (0.69 Å backbone r.m.s.d.), the proline-rich C-terminal “tail” of Cas1a is distinctively ordered in only the bound state and is stabilized by hydrophobic and electrostatic contacts (Fig. 3b). At the middle of the tail, I291 from Cas1 is positioned in a hydrophobic pocket of Cas2 that includes W44 and W60 (Fig. 3b). The C-terminus of Cas1c is likely to span the opposite face of Cas2 to fully encapsulate the dimer, although electron density for this tail was not observed due to crystal packing of an adjacent complex on this face. We also observe structural rearrangements in the C-terminal α-helical domains of Cas1b and Cas1d as compared to apo Cas1 (Fig. 2b), however it is unclear whether this conformational change is due to complex formation or crystallographic packing of another complex next to these monomers.

Cas1–Cas2 complex formation is required in vivo

To determine the function of Cas1–Cas2 complex formation, we conducted spacer acquisition assays in cells expressing Cas1 and Cas2 bearing mutations at the inter-protein interfaces. We found that a structured Cas2 C-terminus is critical for function as its deletion (Δβ6–β7) prevented detectable spacer acquisition (Fig. 3c). This deletion removes the backbone interaction of Cas2 with β4 of Cas1, as well as the D84 residue at the electrostatic interface. In contrast, deletion of the Cas1 tail (ΔP282–S305) did not abolish spacer acquisition. Furthermore, mutation of Cas1 I291, which binds in a hydrophobic pocket with W44 and W60 of Cas2, had no effect in spacer acquisition (Fig. 3a, c). Thus, the C-terminal tail of Cas1 is not essential for spacer acquisition, although it may supplement the critical interactions at the interface described below.

Mutations of residues involved in the electrostatic interactions between subunits have drastic effects on spacer acquisition. Cas1 constructs with mutations at the arginines of α8 to alanine (R245, R252 or R256) could still acquire spacers. However, constructs with mutations of the same residues to the opposite charge (R245D, R252E or R256E) supported little or no detectable spacer acquisition compared to wild-type Cas1 (Fig. 3d). To show that these mutations have little or no effect on Cas1 stability, we purified the R252E mutant to homogeneity and the mutant eluted at the expected retention time for wild-type Cas1 dimer (Supplementary Fig. 2d). In comparison, single mutations of either of the two acidic Cas2 interface residues E65 or D84 to alanine or arginine had little or no effect on spacer acquisition compared to wild-type Cas2. A double mutation of both residues to arginine (E65R and D84R) abolished spacer acquisition in vivo (Fig. 3e). Thus, while mutations in Cas1 at the electrostatic interface are more deleterious than those in Cas2, complementary charges in this interface permit acquisition.

To confirm that the observed in vivo effects are due to disruption of Cas1–Cas2 complex formation, we conducted FLAG immunoprecipitation experiments in lysates of BL21-AI cells overexpressing Cas1-FLAG and Cas2-HA mutants. Mutations that had little effect on spacer acquisition (Cas1 Δtail and Cas2 E65R) did not perturb co-precipitation of Cas1 and Cas2, indicating that the mutants are still able to form the complex (Fig. 3f). In contrast, acquisition-defective mutants (Cas1 R252E and Cas2 Δβ6–β7) can no longer form a stable Cas1–Cas2 complex. This result highlights the importance of the structured Cas2 C-terminus and the positioning of R252 between the two Cas2 acidic residues in the electrostatic interaction interface. Together, these findings support the conclusion that Cas1–Cas2 complex formation is required for spacer acquisition in vivo.

The catalytic activity of Cas2 is dispensable in vivo

Despite the available literature on the biochemical activities of Cas1 and Cas2, the functional roles of these proteins during spacer acquisition are unknown. Cas1 is reported to be a sequence-independent, metal-dependent nuclease that can cleave ssDNA, linear and plasmid dsDNA, ssRNA and various DNA repair intermediates such as Holliday junctions^23,24,26,31. Three different Cas2 homologs were found to have metal-dependent nuclease activity with a preference for ssRNA²² or dsDNA²⁵ or to lack any detectable nuclease activity²⁹. The heterohexameric Cas1-Cas2 complex has five potential active sites-one for each Cas1 monomer and one at the Cas2 homodimer interface (Fig. 4a, b). We conducted spacer acquisition assays with active site residue mutations in Cas1 and Cas2 to determine if the nuclease activities of both proteins are required.

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Figure 4

Complex formation is required for CRISPR DNA recognition

(a,b) Close-up views of the Cas1 and Cas2 active sites with stick representations for the conserved residues mutated in vivo. In the middle is a general view of the active sites in the complex, highlighted in red. (c,d) Acquisition assays of active site residue mutations of Cas1 and Cas2. (e) Western blot of fractions in the biotinylated DNA affinity precipitations. The cartoon representations are the DNA constructs used with the 5′-biotin labels (stars). W.C.L. refers to the whole cell lysate. (f) Western blot of the DNA affinity precipitations in BL21-AI lysates from overexpression of Cas1-FLAG only, Cas2-HA only or both. (g) FLAG IP in lysates from overexpression of Cas1-FLAG and Cas2 E9A-HA. (h) DNA affinity precipitations in the same lysates as in (g) using the same DNA constructs as in (e).

Alanine substitution of the conserved Cas1 active site residues abolished spacer acquisition, demonstrating its critical role in metal-dependent DNA cleavage during the adaptation stage (Fig. 4c). Despite the low protein sequence conservation of Cas2 proteins, a conserved acidic residue from each monomer is positioned in the active site to coordinate a metal ion during catalysis in vitro^22,25 (Fig. 4b). Surprisingly, Cas2 mutated in the signature catalytic E9 residue to alanine or arginine supported spacer acquisition at frequencies similar to those observed in the presence of wild-type Cas2 (Fig. 4d). A mutation of this acidic residue was previously shown to have drastic effects on nucleic acid substrate cleavage in vitro^22,25. Cas2 with an R14A mutation, which was also shown to be catalytically inactive in the Sulfolobus solfataricus Cas2 in vitro²², was still active in acquiring spacers. Of the nearby arginine residues, only the R18A construct had low spacer acquisition levels. This residue interacts with Cas1 at the inter-protein interface, as supported by its continuous electron density with the Cas1 backbone. These results support the notion that Cas1 is the likely nuclease that catalyzes the integration reaction, whereas the function of Cas2 during CRISPR-Cas immunity may not be nucleic acid cleavage.

In vitro complex formation, crystallization and structure determination

Purified Cas1 and Cas2 were separately dialyzed against 150 mM KCl, 20 mM HEPES-KOH, pH 7.4, 5% glycerol and 1 mM TCEP at 4°C overnight. The proteins were incubated together at a 1:3 Cas1:Cas2 molar ratio for one hour on ice. The sample was loaded on a Superdex 75 (16/60) size exclusion column and the peak fractions corresponding to the complex were pooled and concentrated for crystallization. We note that the molar ratio of Cas1:Cas2 for pre-incubation was chosen to obtain clear separation between the Cas1–Cas2 complex and Cas1 only peaks on gel filtration. There is no difference in the retention time of the complex when the proteins were pre-incubated at a 1:1 ratio.

The selenomethionine-derivatized complex was concentrated to ~4 mg mL⁻¹ and crystallized by hanging drop vapor diffusion at room temperature in 120 mM calcium acetate, 10% (w/v) PEG 8000 and 50 mM sodium cacodylate, pH 6.50. The crystals were briefly transferred into a drop containing 20% glycerol for cryoprotection and frozen in liquid nitrogen until data collection. The native protein was crystallized in 150 mM NaCl, 6% (w/v) PEG 8000 and 100 mM Tris, pH 8.0. The crystals were frozen in the presence of 20% PEG 8000 as cryoprotectant.

The diffraction data were collected under cryogenic conditions at the Lawrence Berkeley National Laboratory Advanced Light Source (beamline 8.3.1). The selenomethionine-derivative crystals were scanned for fluorescence to determine the selenium absorption edge and diffraction data was collected at the peak wavelength.. The data were processed with XDS³⁷ and SCALA³⁸. The crystals belonged to a monoclinic space group (P 2₁) with four copies of Cas1 and two copies of Cas2 in the asymmetric unit. Autosol was used for experimental phasing and AutoBuild within PHENIX³⁹ to obtain an initial model. Iterative rounds of model building in Coot⁴⁰ and refinement using PHENIX was performed to obtain a selenomethionine-derivative model. The 2.3-Å native model was built using Phaser–Molecular Replacement in PHENIX for phasing, followed by Coot and PHENIX for model building and refinement, respectively.

Isothermal titration calorimetry

ITC experiments were conducted on a MicroCal Auto-iTC200 system (GE Healthcare). Purified Cas1 and Cas2 were separately dialyzed at 4°C against 200 mM KCl, 20 mM HEPES-KOH, pH 7.5, 5% glycerol and 1 mM TCEP. Cas1 (150 μM) was titrated into the cell containing 15 μM Cas2 with 10–15 3.1 μl injections at 4°C. The Origin software (OriginLab) was used for baseline correction, integration and curve fitting. The K_d reported is an average of three separate experiments (308, 212 and 351 nM, standard deviation of 58 nM). The calculated N values of each independent run were 1.52, 1.51 and 1.50.

Analytical ultracentrifugation

Sedimentation velocity experiments were conducted at 50,000 rpm using the Beckman Coulter XLI (Beckman Coulter, Fullerton, CA, USA). The samples were monitored by absorbance optics at 280 nm. The proteins were dialyzed against 20 mM HEPES-KOH pH 7.5, 150 mM KCl and 1 mM TCEP. Three concentration series for Cas1 at 7, 14 and 28 μM and two concentrations of Cas2 at 20 and 40 μM were conducted to evaluate the formation of higher-order species. The Cas1–Cas2 complex was characterized using a constant concentration of 5 μM of Cas1 and two concentrations of Cas2 at 5 and 10 μM. The solvent density (1.00688 g.ml⁻¹), viscosity (0.01022 poise), and the partial specific volumes that were used for the analyses, 0.7453 ml.g⁻¹ (Cas1), 0.7443 ml.g⁻¹ (Cas2), 0.7451 ml.g⁻¹ (Cas1-Cas2), were calculated by SEDNTERP v. 20120828⁴¹. The sedimentation coefficients and apparent molecular weights were calculated from size distribution analyses [c(s)] using SEDFIT v. 14.3e ^42,43. The figures were prepared using Origin v. 6.0 (Microcal Software Inc.).

Spacer acquisition assays

The assays were conducted as previously described⁵ with slight modifications. Briefly, cas1 and cas2 were both cloned into pCDF-1b (Novagen) and transformed into E. coli BL21-AI (Invitrogen). After preparation of an overnight culture in LB medium containing 50 μg ml⁻¹ streptomycin, a sample was transferred (1:300) into a 10 ml culture containing 0.2% L-arabinose, 0.1 mM IPTG and 50 μg ml⁻¹ streptomycin to induce protein expression. After 20–24 hours, a sample of the culture was diluted in water, boiled at 95°C for 5 min and centrifuged (16,100 × g). A sample of the supernatant was used as template for PCR amplification of the CRISPR locus using the same primers as previously described⁵. The PCR reactions were analyzed on 1.5% agarose gels. For comparison in spacer acquisition efficacy of mutant proteins, the OD₆₀₀ of each culture was measured and the amount of culture obtained for PCR amplification was normalized accordingly. The newly acquired spacers reported in Supplementary Table 1 were obtained by plating a sample of the culture on LB agar plates and amplifying the CRISPR-I locus of single clones to detect locus expansion. The PCR products of clones with expanded loci were submitted for sequencing. The gene annotations were obtained from the NCBI Basic Local Alignment Search Tool (BLAST)⁴⁴ using the Escherichia coli BL21 (taxid:469008) genomic sequence. All of the acquisition assays reported in this study have been replicated at least three times.

Immunoprecipitation assays

The Cas1-FLAG and Cas2-HA constructs were both cloned into pCDF-1b and co-expressed in BL21-AI cells for 20–24 hours in spacer acquisition-inducing conditions, as described above. The amount of cells used was normalized to the OD₆₀₀ measurements of the cultures. The cells were pelleted and re-suspended in lysis buffer (150 mM KCl, 50 mM Tris, pH 7.50, 1 mM TCEP, 1% Triton X-100, 0.5 mM PMSF and protease inhibitors). After sonication on ice, the lysates were cleared and rocked for 1.5–2 hours at 4°C with either anti-FLAG M2 or anti-HA affinity resin (Sigma-Aldrich). The resin was washed five times with 400 mM KCl, 50 mM Tris, pH 7.50, 1 mM TCEP and 1% Triton X-100. The proteins were eluted with either 100 ng μl⁻¹ 3X FLAG peptide (DYKDDDDK) or HA peptide (YPYDVPDYA), synthesized by David King (HHMI, UC Berkeley). The epitope-tagged proteins were detected with monoclonal anti-FLAG M2-peroxidase (HRP) mouse antibody (Sigma-Aldrich A8592, 1:22,000) or HRP-conjugated anti-HA mouse antibody (Cell Signaling 2999S, 1:10,000). All of the FLAG immunoprecipitation experiments reported in this study have been replicated at least three times.

DNA affinity precipitation assays

The 186-bp CRISPR DNA bait was generated by PCR amplification of the BL21-AI CRISPR-I locus using 5′ biotin-conjugated forward and reverse primers (synthesized by Integrated Device Technology). The 186-bp control DNA was PCR amplified from the ori sequence of the pUC19 vector. The input lysates were prepared as described above for IP assays. The amount of biotinylated DNA probe was normalized to 100 nM and rocked with the lysate at 4°C for 30 min. Avidin agarose (Pierce, Fig. 4e) or streptavidin magnetic beads (NEB, Supplementary Fig. 4b) was added to the reaction and rocked for an additional 1.5–2 hours. The samples were washed five times with lysis buffer and the proteins were eluted with Laemmli buffer by boiling at 95°C for 5 minutes. Western blotting was conducted to detect Cas1-FLAG and Cas2-HA in the samples as described in the IP procedure.

We are grateful for the input on this work provided by members of the Doudna lab. We thank S. Floor, A.S. Lee, H.Y. Lee, R. Wilson, R. Wu and K. Zhou for technical assistance, the 8.3.1 beamline staff at the Advanced Light Source and A. Iavarone (UC Berkeley) for mass spectrometry. This project was funded by a US National Science Foundation grant to J.A.D (No. 1244557). J.K.N. and A.V.W. are supported by US National Science Foundation Graduate Research Fellowships and J.K.N. by a UC Berkeley Chancellor’s Fellowship. P.J.K. is a Howard Hughes Medical Institute Fellow of the Life Sciences Research Foundation. J.N. is supported by a Long-Term Postdoctoral Fellowship from the Human Frontier Science Program Organization. J.A.D. is an Investigator of the Howard Hughes Medical Institute.