Synthesis of the IHA ligand

O-tritylhydroxylamine was synthesized as previously described32. Chloroacetyl chloride (0.58 ml, 7.3 mmol) was dissolved in 2 ml CH 2 Cl 2 and added dropwise to a suspension of O-tritylhydroxylamine (2.0 g, 7.3 mmol) and N,N-diisopropylethylamine (2.5 ml, 14.5 mmol) in 15 ml CH 2 Cl 2 at 0 °C. The reaction mixture was gradually warmed to room temperature and stirred at room temperature for an hour. An additional 15 ml CH 2 Cl 2 was added and the reaction was extracted with H 2 O (3 × 30 ml). The CH 2 Cl 2 solution was collected and evaporated to dryness. A solution containing 15 ml of CH 2 Cl 2 with 10% (v/v) trifluoroacetic acid was added and the solution was stirred for 30 min. The crude product was purified by silica gel chromatography using a gradient of 0–100% ethyl acetate in hexanes as the eluent. The product was visualized using a FeCl 3 stain. Yield, 55%. Measured molecular weight (m/z): 108.37 [M − H+]; calculated: 107.99 [M − H+]. 1H NMR: (300 MHz, DMSO-d 6 ) δ 10.88 (s, 1H), δ 9.15 (s, 1H), δ 3.93 (s, 2H). 13C NMR: (500 MHz, DMSO-d 6 ) δ 162.88, δ 40.45. 2-chloro-N-hydroxyacetamide (400 mg, 3.7 mmol) and NaI (2.7 g, 18.3 mmol) were refluxed in 30 ml acetone for 1 h. The reaction mixture was purified by silica gel chromatography with 100% ethyl acetate as the eluent and dried in vacuo. Yield, >90%. Measured molecular weight (m/z): 223.85 [M + Na+]; calculated: 223.95 [M + Na+]. 1H NMR: (300 MHz, DMSO-d 6 ) δ 10.81 (s, 1H), δ 9.09 (s, 1H), δ 3.51 (s, 2H). 13C NMR: (500 MHz, DMSO-d 6 ) δ 164.83, δ −2.01.

Protein expression and purification

All constructs (Supplementary Table 1) were derived from the parent pET-20b(+) plasmid containing the CFMC1 gene via site-directed mutagenesis as previously described28,33,34. The appropriate plasmids were transformed into BL21(DE3) Escherichia coli cells (New England Biolabs) housing a CCM (cytochrome C maturation) cassette containing a chloramphenicol-resistance marker and expressed as previously described35 with minor adjustments. Multiple 2.8-l flasks containing 1.5 l of LB medium were shaken at 200 rpm for 12 h at 37 °C and then at 100 rpm for an additional period of around 7 h. Cells were collected by centrifugation (5,000 rpm for 10 min at 4 °C), resuspended in a buffered solution containing 5 mM sodium acetate (NaOAc) (pH 5.0) and 2 mM dithiothreitol (DTT) and lysed via sonication. The pH of the crude lysate was first raised to 10 using NaOH to precipitate cellular contaminants, then reduced to pH 4.5. After centrifugation (12,000 rpm for 20 min at 4 °C), the clarified supernatant was decanted and diluted 15-fold with additional buffer. This solution was applied to a CM sepharose gravity column (GE Healthcare) pre-equilibrated with the aforementioned buffer and subjected to multiple buffer washes before elution using a stepwise-gradient of NaCl (0–0.5 M). Peak elution fractions were combined and concentrated using a 400-ml Amicon Stirred Cell (Millipore) and buffer-exchanged by overnight dialysis against a buffered solution containing 10 mM phosphate (pH 8.0) at 4 °C. Next, the protein was purified via a DuoFlow workstation station fitted with a Macroprep High Q-cartridge column (BioRad) and eluted using a linear gradient over 0–0.5 M NaCl. Fractions that exhibited an RZ ratio (A 421 /A 280 ) > 4.4 were pooled, treated with 2 mM EDTA for 1 h, concentrated, and buffer-exchanged into 20 mM tris(hydroxymethyl)aminomethane (Tris) (pH 7.5) pretreated with Chelex 100 resin (BioRad), via desalting column (Econo-Pac 10DG pre-packed columns, BioRad). Demetallated and purified proteins were concentrated to around 2 mM and stored at 4 °C.

Protein labelling and post-labelling purification

Purified protein solutions were treated with a 100-fold excess of DTT and placed in an anaerobic Coy chamber for approximately 2 h for slow degassing to remove dissolved oxygen. The fully reduced protein solution was buffer-exchanged into 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.5) via desalting column to remove DTT, and the concentration of the resulting protein solution was determined spectroscopically (Agilent 8452 spectrophotometer) using the ε 421 (red) = 162,000 M−1 cm−1 (ref. 34). Solid iodohydroxamic acid (IHA) was dissolved in 100 μl degassed DMF to generate solutions containing a 15-fold excess IHA per protein monomer, which were then added to protein aliquots and incubated overnight. The HA-functionalized variants were removed from the Coy chamber and separated from unreacted or partially reacted protein via FPLC using a Q-column equilibrated with 10 mM N-cyclohexyl-2-aminoethanesulfonic acid (CHES) (pH 9.3) and 2 mM DTT and eluted using a linear gradient over 0–0.5 M NaCl. Protein functionalization was verified using electrospray ionization mass spectrometry (ESI-MS; Extended Data Fig. 1) and the resulting protein solutions were buffer-exchanged into demetallated 20 mM Tris (pH 7.5) via desalting column, concentrated to around 2 mM and stored at 4 °C for further use.

Redesign of CFMC1 interfaces

To render the CFMC1 protomer competent for the bimetallic design strategy, we first performed the following mutations to remove potential competitive interactions: C67E, H59S and H73N. A negative design strategy was then used to disrupt a noncovalent dimerization interface found in CFMC1, leading to the mutations A34Q and A38Q. We further identified the dearth of protein–protein interactions within the core and periphery of the three-fold axis engulfing the 82 position as a likely contributor to poor cage assembly and crystallization in general. Accordingly, as a means to facilitate cage formation, we adopted Rosetta-prescribed mutations at the following positions: A24T, Q25(T/E), N80K and E81Q.

Crystallography

Screening and crystallization of all BMC variants were conducted via sitting drop vapour diffusion. In brief, solutions containing 2.1–2.2 mM BMC protomer were mixed with mother liquor (1 μl + 1 μl) and equilibrated against 200-μl reservoir volumes. Supplementary Table 2 details the experimental conditions for crystal growth. Protein solutions of BMC1 and BMC4 were first incubated with FeSO 4 for 1 h before mixing with ZnCl 2 . Solutions of BMC2 and BMC3 were mixed with FeSO 4 and ZnCl 2 stock solutions and were immediately combined with the mother liquor (to prevent rapid aggregation of the proteins functionalized with two HA units). Crystals for all mutants typically appeared within several hours and were collected within a week of maturation. Crystals were cryoprotected by submersion into perfluoropolyether cryo oil (Hampton Research) for a few seconds and flash-frozen in liquid nitrogen. X-ray diffraction data were collected at 100 K at either the Advanced Light Source (ALS) beamline BL 8.3.1 (using 1.12 Å radiation for BMC3 and 1.33 Å radiation for BMC4) or at the Stanford Synchrotron Radiation Lightsource (SSRL) beamlines 9-2 (using 0.98 Å radiation for BMC2) and 12-2 (using 0.98 Å radiation for BMC1). Data integration was performed using the XDS Program Package, truncated at CC 1/2 > 0.5 (ref. 36). Datasets of the same structure recorded at different wavelengths were scaled to the highest resolution dataset with XSCALE37,38. Phaser-MR39 was used to carry out molecular replacement with search models based on the CMFC1 monomer (PDB ID: 3M4B) containing the expected side chain mutations (generated in Pymol40) but lacking HA. Rigid-body and structure refinement was performed using multiple rounds of Phenix.refine39, interspersed with manual model rebuilding and metal/ligand placement with Coot41. Restraint files for the Cys-hydroxamic acid conjugates were generated using phenix.eLBOW to maintain the distances Cys-SG–HA-C1 (1.816 Å ± 0.02 Å) and angles Cys-CB–Cys-SG–HA-C1 as well as Cys-SG–HA-C1–HA-C2 (both 109° ± 3°) during refinement. Where necessary, the metal binding geometry of the hydroxamic acids was restrained to the distances Fe–HA-O1 (1.98 Å ± 0.05 Å) and Fe–HA-O2 (2.057 Å ± 0.05 Å) as well as through a planarity constraint for the atoms Fe, HA-O1, HA-O2 and HA-C1 following data from a high-resolution structure of Fe(iii)-tris-benzhydroxamate trihydrate42. Simulated annealing omit maps (metal atoms and side chain ligands) were generated for each metal binding site and model accuracy was assessed critically against these omit maps. Electron density maps were generated using Phenix and all molecular graphics images were produced with either Pymol or the UCSF ChimeraX package from the Computer Graphics Laboratory, University of California, San Francisco43.

Crystallographic metal content analysis

Metal ions, with their relatively high-energy inner electrons, can absorb and resonate with soft X-rays; this leads, among other effects, to differences in the intensity of otherwise centro-symmetric Bragg diffraction peaks used for X-ray crystallography. Density maps calculated from these differences are routinely used to locate and identify metal ions in protein crystals. The magnitude of this anomalous X-ray diffraction varies with the X-ray energy, with stark differences around the energies of the K- and L-shell electrons of the respective elements allowing one to discern between elements at a position in question, if diffraction datasets are measured at the appropriate wavelengths. For a visual analysis of the bound metals, the scaled datasets of different wavelengths were used separately as an input for a single phenix.refine run, each with the final model of the highest resolution dataset. Importantly, only the B-factor or occupancy were allowed to change during refinement, resulting in anomalous difference density maps for each wavelength. Using these maps, isomorphous difference maps from data at wavelengths above and below the respective element K-edges were generated (if applicable) with Phenix and were inspected manually (Extended Data Fig. 5). To gain a more quantitative understanding of the identity of the bound metals for each site, the anomalous difference signal of each dataset was used to generate CCP4 format maps with phenix.mtz2map. The generated maps were used subsequently as inputs to calculate the mean signal in a sphere of 1 Å radius centred on each metal atom with the program MAPMAN (Uppsala Software Factory). For each pair of datasets above and below a metal-absorption edge, the ratio of the anomalous signal above and below the edge for every metal atom was tabulated. The experimental ratio was compared to the theoretical ratio for both Fe and Zn (Extended Data Fig. 5) according to http://skuld.bmsc.washington.edu/scatter, as calculated using the Cromer and Liberman approximation. Theoretical ratios were also calculated for hypothetical mixed occupancy Fe/Zn metal sites and compared to experimentally observed values (Supplementary Tables 3–6).

Protein cage sample preparation

Self-assembled cages

All samples were prepared in a low-O 2 atmosphere (Coy glovebox) to minimize undesired oxidation of Fe2+ ions before self-assembly. Protein solutions containing 20 μM BMC3 or 100 μM BMC4 in 20 mM Tris (pH 8.5) were incubated with either 20 μM FeSO 4 and 60 μM ZnCl 2 for BMC3 or 50 μM FeSO 4 and 200 μM ZnCl 2 for BMC4 for 2–3 h to yield the metallated cages. We note that the addition of FeSO 4 was followed by a small but observable change in the colour of the solution from red to pink, attributed to a shift of the haem Soret band to longer wavelengths, which suggested reduction of the haem by the ferrous ions and generation of ferric ions in close proximity to HA group(s). The final BMC3 solutions were then concentrated sevenfold before overnight incubation to improve the total cage yield. After self-assembly, the resulting solutions were diluted back to their original concentrations with the self-assembly buffer before characterization.

Dissolved crystals

Fe:Zn:BMC1 and Fe:Zn:BMC2 crystals were dissolved using buffer containing 100 mM HEPES (pH 7.5), 200 mM MgCl 2 and 800 μM ZnCl 2 . Mature crystals were removed from their pedestal droplet, briefly submerged in fresh buffer to remove uncrystallized protein and surface-bound precipitates, and transferred into a new sitting drop crystallization well containing 8 μl buffer solution. The crystals were physically crushed with a small metal scalpel and vigorously pipetted until a large portion of the crystals dissolved. Undissolved crystals were removed by centrifugation (10,000 rpm for 5 min at 25 °C), yielding a light-red supernatant and dark-red precipitate.

Negative-stain transmission electron microscopy

A 4-µl droplet of BMC cages (either self-assembled or from dissolved crystals) was deposited onto formvar/carbon-coated Cu grids (Ted Pella) (pretreated by negative-mode glow discharge up to 15 min beforehand) and allowed to bind for 5 min. The grids were then washed with 50 μl MilliQ water, blotted using Whatman filter paper and stained using 2% uranyl acetate solution in water and blotted again. Grids were imaged using a FEI Sphera transmission electron microscope operating at 200 keV, equipped with an LaB 6 filament and a Gatan 4K CCD camera. Micrographs were collected using objective-lens underfocus settings ranging from 250 nm to 2 µm and analysed using Fiji (http://fiji.sc/Fiji).

Oligomerization state determination using AUC

Sedimentation velocity measurements were performed at 41,000 rpm and 25 °C using an XL-1 analytical ultracentrifuge (Beckman Coulter) equipped with an AN-60 Ti rotor. Data processing was performed using Sedfit44 with the following parameters as calculated using SEDNTERP: viscosity: 0.01000 poise, density: 0.9988 g/ml (self-assembled samples) or viscosity: 0.0113191 poise, density: 1.0196 g/ml (dissolved crystals), and a partial specific volume of 0.7313 ml/g for all samples. All reported results correspond to a confidence level of 0.95.

Preparation of samples involving crystal dissolution

Dissolved crystal samples (BMC1 and BMC2), prepared as described above at ambient conditions, were diluted to 350 μl with 10 mM HEPES (pH 7.5), 200 mM MgCl 2 and 800 μM ZnCl 2 . The solution was clarified via brief centrifugation in order to remove crystal debris and the supernatant was placed inside the cells.

Calculation of BMC void volumes

Structures of complete cage assemblies for BMC2, BMC3 and BMC4 were generated via the application of crystallographic symmetry operations to the fully refined asymmetric unit of each construct. These coordinates were recentred at the origin and stripped of waters, hydrogens, alternative conformations and crystallization reagents (PEG-400). Volumetric maps and volumes for the internal cavity of each cage were calculated using VOIDOO45, and are reported as the solvent-accessible volume for a 1.4 Å rolling probe on a 0.25 Å grid spacing for all constructs. The cavity volumes using these parameters were determined to be approximately 32,700 Å3 (BMC2), 32,700 Å3 (BMC3) and 7,900 Å3 (BMC4).

Solution self-assembly, disassembly and thermal stability of BMC3 and BMC4

Assembled samples were prepared as described above and placed inside the AUC measurement cells anaerobically (20 μM BMC3 and 100 μM BMC4). Disassembly of the cages via metal-ion removal was performed by treating the protein cages with 2 mM EDTA for 1 h. Redox-controlled disassembly of the protein commenced by the addition of either 5 mM sodium dithionite or 5 mM sodium ascorbate to the cage solution anaerobically and subsequent incubation of the samples at around 22 °C for 16 h. Samples were then loaded into the AUC measurement cell.

For thermal stability measurements, samples were placed in a thermoregulated chamber preequilibrated at the appropriate temperature for 2 h, and subsequently removed from the chamber and equilibrated at room temperature for 30 min before AUC analysis. Circular dichroism (CD) spectra were measured using an Aviv 215 spectrometer. CD measurements were performed using 10 µM protein in a buffered solution containing 20 mM Tris (pH 8.5). Thermal melts were measured at 222 nm at a 1 nm slit width, scanning at 1-nm intervals with a 1-s integration time. Measurements were taken from 25 °C to 85 °C at 2-degree intervals with a 2 min equilibration at each temperature. Unfolding data were fit to a two-state model with van’t Hoff’s enthalpy using the CalFitter web server46.

Cryo-EM sample preparation

Self-assembled BMC3 cages were removed from the anaerobic Coy chamber immediately before grid preparation. A 3.5-µl aliquot of self-assembled BMC3 cages was dropped onto holey carbon grids (Electron Microscopy Sciences, Quantifoil R1.2/1.3 holey carbon on 300 mesh copper) that had been freshly glow-discharged for 30 s. The initial application of the sample was side blotted manually with Whatman No. 1 filter paper immediately followed by a secondary application of a 3.5-µl aliquot, blotted for 3.5 s and plunge-frozen in liquid ethane cooled by liquid nitrogen using a Vitrobot Mark IV (FEI).

Cryo-EM data acquisition and image processing

Samples were imaged on a Titan Krios G3 transmission electron microscope (FEI) operating at 300 kV equipped with a K2 Summit direct electron detector (Gatan) and a GIF Quantum energy filter. The slit-width of the energy filter was set to 10 eV. Movies were collected at a magnification of 165,000× in EFTEM mode giving a physical pixel size of 0.84 Å/pixel. In total, 4,672 movie stacks (50 frames/movie) were collected using a 10 s exposure at a dose rate of 1.2 e−/Å2 per frame for a total electron dose of 60 e−/Å2 per movie. Objective-lens underfocus settings varied between 0.6 µm and 1.6 µm. Data collection was performed using software EPU (FEI). All image processing was performed in the Relion-3.0 pipeline47. Motion correction and dose weighting were performed using MotionCor248, and defocus values were estimated with Gctf49 using a pixel size of 0.8 Å/pixel. A total of 3,513 movie stacks were selected following motion correction and CTF estimation, and 805,156 particles were auto-picked using RELION-3.0. Particle images were extracted and binned by 2 (1.6 Å/pixel, 100 pixel box size) and subjected to two-dimensional (2D) classification. A total of 444,247 particles were selected corresponding to good 2D class averages and subjected to three-dimensional (3D) classification imposing T symmetry and using an initial model generated from a subset of the particles. A total of 129,653 particles were chosen from a 3D class showing strong secondary-structural elements and subjected to 3D auto-refinement with T symmetry. The particles were re-centred and re-extracted to their original pixel size of 0.8 Å/pixel. These particles were subjected to 3D auto-refinement with T symmetry and the yield map was then postprocessed towards 2.6 Å resolution based on the gold-standard Fourier shell correlation (FSC) 0.143 criterion. The pixel size of the map was manually adjusted using Relion image handler to match the physical pixel size of the images. Local resolution was calculated in Relion 3.0 using ResMap50.

Model building and refinement

The BMC3 crystal structure (PDB ID: 6OT7) stripped of hydrogens and waters was used as an initial model and manually docked into the cryo-EM density using UCSF Chimera51. The structural model was subject to real space refinement in Phenix against the cryo-EM map with geometry restraints for the Fe-binding sites and molecular coordinates for the Cys–HA ligand. The atomic model was manually improved using Coot. Tightly bound waters were identified based on clear density in the EM density map. Whereas the structural flexibility of the hydroxamate sites manifested in poor electron density, the twofold interface was much more rigid and unambiguous density was observed for Zn-binding. A tryptophan at the 66 position, which had shown high-temperature factors in the BMC3 crystal structure, was identified in multiple conformations in the EM density map. The final model was subjected to real space refinement using Phenix39 and evaluated using MolProbity52. All molecular graphics images were rendered in PyMoL or UCSF ChimeraX.

Encapsulation of rhodamine in BMC3 cages

BMC3 cages were self-assembled in a low-O 2 atmosphere in the presence of rhodamine for the passive encapsulation of the dye. Solutions containing 20 μM BMC3 were incubated with 20 μM FeSO 4 , 60 μM ZnCl 2 and 2 mM rhodamine. A control sample was prepared in the absence of added metal ions (20 μM BMC3 incubated with 2 mM rhodamine). Samples were incubated for 2–3 h and concentrated sevenfold before overnight incubation. Protein solutions were buffer exchanged on a PD-10 desalting column using a buffer containing 20 mM Tris (pH 8.5) (with 5 μM FeSO 4 and 10 μM ZnCl 2 supplemented for solutions already containing metal ions) to separate unassociated dye from protein. Cage solutions were split in two: one half was treated with 1 mM EDTA and incubated for 2 h before washing. All protein solutions were additionally washed three times using a centrifugal filter to completely remove any remaining free rhodamine.

Fluorescence measurements were performed using 6 μM protein solutions after the previously mentioned wash steps. For each sample, an excitation wavelength of 555 nm with a 2 nm slit width was used and emission was measured between 560 and 650 nm with a 2 nm slit width and 0.2 s integration time. For the time-course experiments, cages encapsulating rhodamine were washed three times after 4 days and after 7 days and diluted to 6 μM before fluorescence measurements. AUC measurements were performed at the λ max of the cytochrome (415 nm) and at the λ max of rhodamine (555 nm) to assess whether there was a sufficiently large rhodamine signal associated with BMC3 cages. Ultraviolet–visible light (UV-vis) absorbance measurements were performed on each solution to measure the protein and rhodamine concentrations. Difference spectra were taken between each rhodamine-incubated sample and BMC3 protomer to eliminate any background signal.

Statistics and reproducibility

All reported samples represent technical replicates. The ns-TEM micrograph of BMC2 cages after 3D crystal dissolution (Fig. 2a) is representative of experiments repeated independently four times. AUC experiments for BMC2 (Fig. 2b) were performed in duplicate. Self-assembly of BMC3 cages and subsequent AUC characterization (Fig. 3a) were performed the following number of times: BMC3 protomer (n = 2), +Fe2+ (n = 4), +Zn2+ (n = 4), +Fe2, +Zn2+ (n = 6). Self-assembly of BMC4 cages and subsequent AUC characterization (Fig. 4a) was performed the following number of times: BMC4 protomer (n = 2), +Fe2+ (n = 1), +Zn2+ (n = 1), +Fe2, +Zn2+ (n = 5). Mass spectra (Extended Data Fig. 1c–f) were collected in duplicate for native and HA-labelled proteins; AUC experiments were performed in duplicate. TEM characterization of BMC constructs (Extended Data Fig. 3) were performed the following number of times: dissolved BMC1 crystals (n = 1), dissolved BMC2 crystals (n = 4), BMC2 +EDTA (n = 2), self-assembled BMC3 cages (n = 5), BMC3 +EDTA (n = 4). AUC experiments following the incubation of BMC3 with first-row transition metals (Extended Data Fig. 6a) were performed in duplicate. Self-assembly of BMC3 in the presence of Fe(acetylacetonate) 3 (Extended Data Fig. 6b) was performed in duplicate. BMC3 cage disassembly in the presence of EDTA (Extended Data Fig. 6c) was performed in triplicate. AUC characterization of BMC variants after equilibration at different temperatures (Extended Data Fig. 6d) was performed the following number of times: BMC3 at 50 °C (n = 2), BMC3 at 70 °C (n = 2), BMC4 at 50 °C (n = 3), BMC4 at 70 °C (n = 3), BMC4 at 90 °C (n = 3). Thermal unfolding of BMC variants as measured by CD spectroscopy (Extended Data Fig. 6d) was performed in duplicate. Treatment of BMC3 cages with chemical reductants (Extended Data Fig. 6e) was performed in duplicate. Cryo-EM characterization of BMC3 cages was performed after collecting 4,672 movie stacks. Extended Data Figure 7a shows a representative micrograph and three representative 2D class averages. Fluorescence characterization of BMC3 samples incubated with rhodamine were performed (Extended Data Fig. 8a) in triplicate. AUC characterization of BMC3 cages encapsulating rhodamine (Extended Data Fig. 8b) was performed in duplicate. UV-vis characterization of BMC3 samples incubated with rhodamine (Extended Data Fig. 8c, d) was performed in triplicate. Repeated fluorescence characterization of a solution containing BMC3 cages encapsulating rhodamine (Extended Data Fig. 8e) was performed in duplicate.

Reporting summary

Further information on research design is available in the Nature Research Reporting Summary linked to this paper.