Supercharged proteins and polypeptides for advanced materials in chemistry and biology
Ma, Chao
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113
Chapter 4
Self-Assembly of Electrostatic Cocrystals from
Supercharged Fusion Peptides and Protein Cages
This chapter has been published:
Antti Korpi, Chao Ma, Kai Liu, Nonappa, Andreas Herrmann, Olli Ikkala, and Mauri A. Kostiainen
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Self-assembly is a convenient process to arrange complex biomolecules into large hierarchically ordered structures. Electrostatic attraction between the building blocks is a particularly interesting driving force for the assembly process, as it is easily tunable and reversible. Large biomolecules with high surface charge density, such as proteins and protein cages, are very promising building blocks due to their uniform size and shape. Assemblies of functional molecules with well-defined nanostructures have wide-ranging applications but are difficult to produce precisely by synthetic methods. Furthermore, obtaining highly ordered structures is an important prerequisite for X-ray structure analysis. In this chapter we show how negatively charged ferritin and viral protein cages can adopt specific cocrystal structures with supercharged cationic polypeptides (SUPs, K72) and their recombinant fusions with green fluorescent protein (GFP-K72). The cage structures and recombinant proteins self-assemble in aqueous solution to large ordered structures, where the structure morphology and size are controlled by the ratio of oppositely charged building blocks and the electrolyte concentration. Both ferritin and viral cages form cocrystals with face centered cubic structure and lattice constants of 14.0 and 28.5 nm, respectively. The crystals are porous and the cationic recombinant proteins occupy the voids between the cages. Such systems resemble naturally occurring occlusion bodies and may serve as protecting agents as well as aid the structure determination of biomolecules by X-ray scattering.
M.A.K., A.H. and O.I. conceived the idea. C.M. and K.L. fabricated and expressed the supercharged proteins. A. K. performed assembly characterization and analyzed data. The introduction, results and discussion parts dealing with SUPs were written by C.M. and corrected by A.H..
115 1. INTRODUCTION
Mimicking the highly evolved functionalities of native biomolecules has been in the focus of research efforts, especially over the past decade.[1] Besides chemical composition, functionalities of natural systems are typically based on the three-dimensional position of the molecules. Additionally, biomolecules are often large but can still adopt specific hierarchical structures with great selectivity. Production of synthetic materials that could achieve the same level of structural sophistication has, however, been challenging.[2]
Another way to harvest the designs of nature is to extract the molecules from natural sources and incorporate them into nanostructured materials.[3] The restrictions of top-down methods to produce fine-structures can simultaneously be overcome, as many biological molecules form organized systems via self-assembly processes.[4] The procedure is the basis of many natural phenomena like protein folding[5] and can be used to produce functional materials with well-defined nanostructures.[6] Self-assembly is typically carried out in liquid media, which allows the building blocks to diffuse without restraints.[7] Noncovalent self-assembly is preferred in many cases as it is typically reversible, easy to control, and applicable to a large pool of molecules, allowing production of assemblies with varying chemical composition and physical dimensions.[8] The assemblies can additionally be tuned by chemical modification of the assembling particles or changing the environment of the assembly. Several bottom-up synthesis methods have been recently studied to produce such nanostructured materials.[9−15] Practical applications include tissue engineering,[16] drug delivery,[17,18] catalysis,[19] and nanopatterning.[20]
Protein cages have been utilized as part of self-assembling systems due to their ability to retain functionality while complexed.[21−25] They often possess uniform size and shape, making them ideal building blocks for crystalline assemblies.[24,26] Many protein cages additionally carry an overall electric charge,[27,28] which enables them to assemble via electrostatic interactions. Such assemblies are reversible and responsive to changes in both pH and salinity of the solution, allowing additional control over the system.[29−31] To form complexes, the charged particles require counterparts with opposing charge. Polyelectrolytes are a noteworthy option as they possess high charge density.[32] They also have the ability to provide proteins
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and enzymes with additional stability and have therefore been used in delivery systems.[33] Copolymers enable even more possibilities for optimizing such systems, as block copolymers composed of oppositely charged blocks have been reported to further enhance the stability of protein complexes.[34,35] However, this method restrains the amount of protein binding with the polymers as only a part of the polymer chains can interact with the protein. Introducing additional charges onto the particles or initially selecting proteins with higher charge densities has been found to be an effective way to increase system stability.[36]
We have previously shown that positively charged avidin proteins and negatively charged protein cages can form ordered structures through electrostatic self-assembly.[30] These structures could be further functionalized with different biotintagged moieties. However, this approach requires an additional biotinylation step. To overcome this, we wanted to study whether fusion proteins that are directly produced with a cationic peptide could be incorporated into the crystals structures.
2. RESULTS AND DISCUSSION
In this chapter, we focus on the self-assembly properties of two native protein cages: apoferritin from Pyrococcus furiosus (aFT) and cowpea chlorotic mottle virus (CCMV) from Vigna unguiculata. The two cages are complexed with cationic supercharged polypeptides (SUPs) composed of 72 consecutive lysine-containing repeating units (K72) as well as green fluorescent protein (GFP) produced as a recombinant fusion with the same SUP tag (GFP-K72). The SUPs are derived from elastin-like polypeptides, consisting of pentapeptide repeats (GVGXP) where the fourth position X was substituted with a lysine (K) residue by molecular cloning.[37,38] The structure of the building blocks, including approximate dimensions and the electrostatic surface potential of the cages are presented in Figure 1. The oppositely charged systems were found to self-assemble in aqueous solution at zero or minor electrolyte concentration, but an excessive addition of electrolyte caused the particles to disassemble back into individual molecules, as expected for electrostatically interacting systems.[39]
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Figure 1. Building blocks used in the study. Negatively charged a) CCMV (pI ~3.8)
and b) aFT (pI ~4.5). Calculated crude vacuum electrostatic potential of the full cages (upper) and solution electrostatic surface potential of protein trimer subunits (lower) are presented for both cages. Red and blue colors represent negative and positive electrostatic potential, respectively. Values range from 0 kBT e
-1
(blue) to -9 kBT e
-1
(red), where kB is Boltzmann constant, T absolute temperature and e
elementary charge. c) GFP-K72 and the chemical structure of K72. Both K72 homopolymer and recombinant GFP-K72 were used in this study.
The size and structure morphology of the assemblies were studied and most of them were crystalline with face centered cubic (fcc) morphology. The presence of GFP appeared to hinder the formation of crystalline assemblies, especially in the case of small aFT cage. It should also be noted that a variety of materials (protein, nanoparticle, synthetic small molecule, etc.) that we have tried to coassemble with protein cages, fail to give ordered structures even after thorough optimization. This provides additional support for the benefits of the SUPs studied in this work. The self-assembly process was first studied using dynamic light scattering (DLS) by titrating aqueous aFT or CCMV solution with K72 or GFP-K72. The formation of the assemblies was followed by monitoring the scattering count rate and the hydrodynamic diameter (Dh). In the case of CCMV, the count rate increases
together with the amount of added K72 or GFP-K72 and reached a plateau when cpc
CCMV−1 > 0.5, indicating the formation of large assemblies in the solution (Figure 2a). Count rate did not decrease even if titration was continued further. With aFT, a distinct difference in the count rate behavior was observed. The count rate increased first to high values, after which it descended quickly until it reached and maintained a constant level when cpc aFT−1 > 0.55 (Figure 2b). This indicates the system in
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scattering. The small assemblies merge once they pass a critical concentration and the count rate drops even though the size of the assemblies is constantly growing.[40] The formed structures were disassembled by titration with aqueous sodium chloride (NaCl) solution, as a sufficiently high electrolyte concentration screened the electrostatic interactions between K72 and the protein cages. Both CCMV complexes disassembled uniformly when titrated with NaCl solution (Figure 2c). In the case aFT, a steady decrease in the count rate was observed for K72 complex, but aFT−GFP-K72 underwent an increase in the count rate at the beginning of the NaCl titration (Figure 2d). This indicates that the latter structures did not disassemble uniformly throughout the solution but broke first into numerous smaller assemblies. These small particles also disassembled when cNaCl > 100 mM, and
the count rate of the system settled to approximately the same values that were measured from the aqueous solution of free aFT.
Figure 2. DLS and agarose gel EMSA data. a) CCMV solution (50 mg L-1) titrated with K72 and GFP-K72. b) aFT solution (100 mg L-1) titrated with K72 and GFP-K72. c) Electrolyte induced disassembly of the CCMV complexes. d) Electrolyte
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(NaCl) induced disassembly of the aFT complexes. Volume-average size distribution profiles of CCMV with e) K72, f) GFP-K72 and aFT with g) K72, h) GFP-K72 at different stages of titration (panels a) and b), respectively). i) Agarose gel EMSA demonstrating the effect of increasing K72 and GFP-K72 concentration on the electrophoretic mobility of CCMV.
Dh of the complexes was monitored throughout the K72 and GFP-K72 titrations to
follow the increase in particle size (Figure 2e−h). The complexes were studied (1) at the beginning of the titration, (2) at the concentration where count rate peaked, and (3) at the concentration where the count rate leveled. For CCMV complexes, neither K72 nor GFP-K72 complex grew significantly when using an excess of the protein (Figure 2e,f), suggesting they did not undergo a step with a large number of small particles as aFT complexes did. With aFT, both K72 and GFP-K72 complex kept increasing in size throughout the titration, confirming the hypothesis that the mid titration sharp increase in count rate was due to the amount of the particles, not their size (Figure 2g,h). The final Dh of all four assemblies was close to 1 μm.
The self-assembly of CCMV with K72 and GFP-K72 was further demonstrated using agarose gel electrophoresis mobility shift assay (EMSA). CCMV was complexed with increasing concentrations of the cationic species, causing a loss in electrophoretic mobility as larger assemblies were formed. This was indicated by a tail, which followed the main band containing the smaller and more mobile particles (Figure 2i). The assemblies lost all mobility as the polycation concentration was increased high enough. GFP-K72 complex lost its mobility in lower concentrations than their K72 counterparts, which is in good agreement with the DLS measurements. Zeta potential measurements were conducted to investigate the surface charge of the assemblies. None of the complexes presented significant electrophoretic mobility, indicating surface charge close to zero (Figure S4). The morphology of the formed assemblies was studied using small-angle X-ray scattering (SAXS). The measurements were conducted in 10 mM NaCl solutions. The measured curves for CCMV complexes with both K72 and GFP-K72 (Figure 3a) as well as aFT−K72 complex (Figure 3b) implicated crystalline structures with fcc packing (space group Fm3̅m; number 225, (hkl) = (111), (200), (220), (311), (420), (422); q/q*= 1, √(4/3), √(8/3), √(11/3)). Face-centered cubic (fcc) structures are typical for aFT systems,[41] but CCMV has been reported to adopt both fcc[42] and body-centered cubic (bcc) configurations.[30] aFT complexed with GFP-K72
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was not crystalline, but broad signals were detected at the regions where aFT−K72 showed narrow well-resolved peaks. The assemblies were mostly amorphous and the broad signals were caused by weakly ordered regions. The different morphology explains the difference in the DLS curves between aFT complexes of K72 and GFP-K72, when the complexes were disassembled with NaCl. The amorphous structure of the aFT−GFP-K72 complex is most likely due to the size mismatch of the building blocks. The size of GFP-K72 is too large to fit into the voids between fcc packet aFT particles, which hinders the formation of an ordered structure.
Lattice constants of the K72 complexes with both aFT and CCMV were calculated using a linear fit to the peak positions obtained by SAXS plotted against the quadratic Miller indices of assigned reflections (Figure 3c). The lattice constant (a) for a cubic lattice can be obtained through equation a = 2π√(h2 + k2 + l2)/q(hkl) and
was calculated to be 40.3 for CCMV−K72 and 19.8 nm for aFT−K72. By using these values, the center-tocenter distance (dcc) of both complexes was calculated by
using the equation dcc = a/√2. For CCMV−K72, dcc was 28.5 nm and for aFT−K72
14.0 nm (Figure 3d). These values correspond well with the sizes of aFT and CCMV.
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Figure 3. Structure morphology characterization by SAXS. SAXS profiles of a)
CCMV–K72 and CCMV–GFP-K72 complexes and free CCMV, b) aFT–K72 and aFT–GFP-K72 complexes and free aFT. c) Quadratic Miller indices of assigned reflections for fcc structures versus measured q-vector positions for the indexed peaks. Solid lines present the linear fits. d) Unit cells and dimension details of aFT– K72 (top) and CCMV–GFP-K72 (bottom) crystals. TEM images of the structure morphology of the studied complexes. e) CCM–K72 crystals. f) CCMV–GFP-K72 crystals. Inset shows an optical microscopy image of the complexes. g) CCMV– K72 crystals. h) aFT–GFP-K72 in amorphous state. The magnification shows the well-ordered and amorphous structures of the complexes respectively.
Cryogenic transmission electron microscopy (cryo-TEM) was used to image the nanostructure of the assemblies. As seen in Figure 3, CCMV formed crystalline lattices with both K72 (Figure 3e) and GFP-K72 (Figure 3f). This is evident from the spherical shape and beveled pattern of the assemblies. The individual virus particles, which are ordered into small crystallites, can also be clearly observed. The observed crystallite size varied from approximately 300 nm to 1 μm. As DLS and SAXS measurements suggested, aFT−K72 complexes were crystalline (Figure 3g), whereas aFT−GFPK72 complexes were amorphous (Figure 3h) and lacked both a distinguishable shape and the beveled pattern.
At optimized conditions, the particles could reach diameters large enough to be imaged using optical microscopy (Figure 3f, inset). The effect of electrolyte concentration on the formed assemblies was studied by preparing salinity series from 0 to 150 mM NaCl in which K72 and GFP-K72 were left to form assemblies with both aFT and CCMV over the course of 10 days at 6 °C. K72 complexes with both aFT and CCMV were the largest ones observed and had a diameter from 30 to 100 μm. The assemblies were heavily branched and irregularly shaped crystals. aFT−GFP-K72 complex had varying shape and size, as expected due to its amorphous morphology. CCMV−GFP-K72 complex formed the clearest crystalline structures. All of the large structures could be disassembled by 100 mM NaCl concentration.
Occlusion body mimicking protection of the complexed GFP was studied using fluorescence spectroscopy. Trypsin, an effective protease, was introduced into solutions of GFP-K72 with CCMV or aFT, and quenching of the fluorescence of
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GFP was investigated. Without the presence of CCMV or aFT, the fluorescence decreased as trypsin digested the GFP, but in the complexes this was not observed (Figure S6). For crystalline CCMV-GFP-K72 systems this was to be expected, as the GFP moieties were likely to be contained within the crystal lattices, and the used trypsin concentration could not efficiently digest the outer layers of the assemblies within the time frame of the measurement. Interestingly, fluorescence spectroscopy measurements also suggest that the GFP moieties are sufficiently safeguarded in the aFT complexes as well, regardless of the amorphous nature of the systems.
3. CONCLUSION
In conclusion, K72 and GFP-K72 underwent reversible selfassembly in aqueous solutions with both CCMV and aFT via electrostatic interactions. Electrolyte concentration that exceeded a critical point (~100 mM NaCl) screened the interactions and caused the structures to disassemble. The same effect can most likely be achieved by adjusting the pH of the solution. The complexes adopted fcc packed crystalline morphology except for the aFT−GFP-K72 complex, which had an amorphous structure. This is most likely due to steric hindrance caused by GFP and is not present in CCMV complex because the cavities between the protein cages are large enough to house GFP. All of the complexes assembled into macroscopic structures, demonstrating that additional functionalities can be embedded into the systems without preventing self-assembly. Such structures resemble occlusion bodies found in nature[43] and could find potential applications for maintaining the long-term stability of delicate biomolecules.[44] To make a better comparison of cocrystal assembly using SUPs compared to synthetic polyelectrolytes, a table is prepared in Table S5.
4. EXPERIMENTAL PART AND SUPPLEMENTARY DATA Materials
CCMV particles were grown and isolated from California black-eye beans as reported previously.[45, 46] Apoferritin from Pyrococcus furiosus (aFT) in double distilled water was provided by MoLiRom (www.molirom.com or MoLiRom srl,
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via Ravenna 8, 00161 Rome, Italy). NaCl (≥ 99.8 %) was ordered from Sigma-Aldrich and used as received. Agarose (sulphate ≤ 0.08 %) was ordered from Biotop and used as received. 50X TAE buffer was ordered from Omega Bio-tek and used as received. All water used was Milli-Q quality.
Protein expression and purification
The construction of cloning and expression vectors were performed as reported from our group.[37, 38] In brief, the SUP genes in the cloning vector were synthesized via recursive directional ligation[47], afterwards the gene fragments were subcloned into the expression vector pET25b(+). The amino acids sequences of K72 and GFP-K72 are shown below in Figure S1.
E.coli BLR (DE3) competent cells (Novagen) were transformed with the expression vectors containing protein of interests. Terrific Broth medium (for 1 L, 12 g tryptone and 24 g yeast extract) enriched with phosphate buffer (for 1 L, 2.31 g potassium phosphate monobasic and 12.54 g potassium phosphate dibasic) and glycerol (4 mL per liter TB) and supplemented with 100 µg/mL ampicillin, was inoculated with an overnight starter culture to an initial optical density at 600 nm (OD600) of 0.1 and incubated at 37°C with orbital agitation at 250 rpm until OD600 reached 0.7. Protein expression was induced by shifting temperature to 30°C. Cultures were then continued for additional 16h post-induction. Cells were subsequently harvested by centrifugation (7,000 x g, 30 min, 4ºC), resuspended in lysis buffer (50 mM sodium phosphate buffer, pH 8.0, 300 mM NaCl, 20 mM imidazole) to an OD600 of 100 and disrupted with a constant cell disrupter (Constant Systems Ltd., UK). Cell debris was removed by centrifugation (40,000 x g, 90 min, 4ºC). Proteins were purified from the supernatant under native conditions by Ni-sepharose chromatography. Product-containing fractions were pooled and dialyzed against ultrapure water and then purified by anion exchange chromatography using a Q HP column. Protein-containing fractions were dialyzed extensively against ultrapure water. Purified proteins were frozen in liquid nitrogen, lyophilized and stored at -80ºC until further use.
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Figure S1. The amino acid sequences of K72(a) and GFP-K72(b) used in this
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Protein Characterization with SDS-PAGE and Mass Spectrometry
The concentrations of the purified polypeptides were determined by measuring absorbance at 280 nm using a spectrophotometer (Spectra Max M2, Molecular Devices, Sunnyvale, USA). Product purity was determined by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) on a 15% polyacrylamide gel. Afterwards, gels were stained with Coomassie staining solution (40% methanol, 10% glacial acetic acid, 1 g/L Brilliant Blue R250). Photographs of the gels after staining were taken with a LAS-3000 Image Reader (Fuji Photo Film GmbH, Dusseldorf, Germany). The result is shown as Figure S2, where the supercharged polypeptides exhibit different electrophoretic mobility.
Figure S2. SDS-PAGE result of K72 and K72. Lane 1, K72. Lane 2,
GFP-K72. M, protein ladder. The electrophoresis behavior of those proteins are different from neutral proteins because of excess amount of charges.
Mass spectrometric analysis was performed using a 4800 MALDI-TOF Analyzer in the linear positive mode. The protein samples were mixed 1:1 v/v with α-Cyano-4-hydroxycinnamic acid (SIGMA) (100 mg/mL in 70% ACN and 0.1% TFA). Mass spectra were analyzed with the Data Explorer software (version 4.9). Values determined by mass spectrometry are in good agreement with the masses that are
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calculated (shown in Figure S3 and Table S1) based on the amino acid sequence.
Figure S3. MALDI-TOF mass spectra of proteins used in this study. Table S1. Mass determination of GFP-ELP variants.
*average molecular weight calculated with ProtParam tool.
#molecular weight determined by MALDI-TOF mass spectrometry.
Dynamic Light Scattering (DLS) and Zeta Potential
Scattering intensity and hydrodynamic radius of the studied complexes were measured using a Nano ZS ZEN3600 tabletop DLS device provided by Malvern Instruments. The same instrument was used for the zeta potential measurements. All measurements were carried out at room temperature using water as solvent. The
M calc* [Da] M ms# [Da]
K72 36445 36426+/-50
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initial samples in DLS measurements were prepared by adding 10 µL of CCMV stock solution (1 g/L) into 490 µL of water or 5 µL of aFT stock solution (10 g/L in water) into 495 µL of water, making the concentration of CCMV solution 0.02 g/L and that of aFT solution 0.1 g/L. K72 or GFP-K72 were gradually titrated into the initial solutions until the count rate stabilized. As the concentrations of CCMV and aFT differed from each other, the concentration of K72 and GFP-K72 were announced as concentration ratio to CCMV or aFT to make comparison easier. The solution was mixed with a micropipette after each addition and before measuring. Brand semi-micro UV-Cuvettes (PMMA) were used for all measurements. Count rates and volume distributions were obtained using Zetaziser software by Malvern Instruments. The volume distribution curves were presented for the initial solution, the end of steep increase in count rate and the final stabilized state (denoted as 1, 2 and 3 in Figure 2, respectively). Afterwards, the solutions were titrated with 5 M NaCl solution in water to investigate the disassembly of the structures. The titration was carried out similarly as K72 and GFP-K72 titrations, gradually and with mixing after each titration step.
In zeta potential measurements the CCMV and aFT water solutions were prepared in a similar manner to the DLS samples, making the concentration of the CCMV solution 0.02 g/L and that of aFT solutions 0.1g/L in water. 6 µL of K72 or GFP-K72 stock solution (1 g/L) were added to the CCMV solutions and 30µL of the same K72 or GFP-K72 stock solutions to the aFT solutions. After the zeta potentials of these final solutions were measured, the K72 or GFP-K72 content was doubled by adding another 6 µL of the stock solutions to the CCMV samples and 30 µL to the aFT solutions in order to see if excessive cation concentration would have an impact on the measurements. The zeta potential curves are presented in Figure S4.
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Figure S4. Zeta potential curves with ratio of K72 or GFP-K72 to CCMV or aFT a)
3:5 and b) 6:5.
Agarose Gel Electrophoresis Mobility Shift Assay (EMSA)
Electrophoresis mobility of CCMV was followed as a function of K72 and GFP-K72 concentrations. The gel was prepared by dissolving 1 g of agarose into 100 mL
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of 1 x TAE buffer in a microwave oven. The solution was stained with 100 µL of 0.625 g/L ethidium bromide, poured into a mold and allowed to set while cooling down to room temperature. Samples were prepared by mixing CCMV and GFP-K72 solutions in water as presented in Table S2.
Table S2. CCMV samples for agarose gel EMSA.
1 µL of 6X blue gel loading dye was added into each sample and they were gently mixed with a pipette. 8 µL of each solution was inserted into separate wells in the gels. The gel was run at 145 V for 30 minutes with 10 mM NaAc (pH 4.8) as the running buffer. Gel was imaged using Bio-Rad Gel DocTM EZ imaging system.
Small-angle X-ray Scattering (SAXS)
Scattering was measured using a system consisting of rotating anode Bruker Microstar microfocus X-raysource (Cu Kα radiation, λ = 1.54 Å), Montel multilayer focusing monochromator (Incoatec), four collimating slits (JJ X-Ray, resulting in beam size of less than 1 mm at the sample position) and Hi-Star 2D area detector (Bruker, sample to detector distance 1.59 m). The instrumentation except for the detector were under high vacuum to prevent scattering from the air. Silver behenate standard was used for the calibration of the scattering vector 𝑞 (magnitude of 𝑞 = 4𝜋𝑠𝑖𝑛𝜃/𝜆, where 2𝜃 is the scattering angle) and the onedimensional SAXS data was obtained by azimuthally averaging the 2D scattering data. The samples were prepared by mixing together 1.0 µL of K72 or GFP-K72, 4 µL of water and 5 µL of CCMV stock solution (1 g/L) or 2.5 µL of K72 or GFP-K72, 2.5 µL water and 5.0 µL of aFT stock solution (1 g/L). Thus the weight ratios were 1:0.2 for CCMV
130
samples and 1:0.5 for aFT samples. The sample solutions were sealed within a steel slug sealed from both sides with Kapton foil.
Cryo-Transmission Electron Microscopy (Cryo-TEM)
The transmission electron microscopy (TEM) images were collected using JEM 3200FSC field emission microscope (JEOL) operated at 300 kV in bright field mode with Omega-type Zero-loss energy filter. Samples were imaged on plasma cleaned 200-mesh copper grids with either holey carbon (CF-Quantifoil) or lacey carbon support film. 3.0 µL of the aqueous dispersion was placed on a grid and plunge freezed in 1/1 (v/v) liquid propane/ethane mixture using Vitrobot™ with 3s blotting time under 100 % humidity. The images were acquired with GATAN DIGITAL MICROGRAPH software while the specimen temperature was maintained at -187 °C.
Optical Microscopy
The optical microscopy imaging was done using Leica DM4500 P microscope combined with Canon EOS60D camera in transmission mode. Samples were prepared in tris(hydroxymethyl)aminomethane (Tris) buffer with varying NaCl concentration to observe the effect of electrolyte. The samples were prepared by making a Tris buffer in water and adding NaCl, K72 or GFP-K72 and CCMV or aFT into it (in that order) and mixing gently with a pipette. All of the solutions had 5.0 mM of Tris and 100 mg/L of aFT or 50 mg/L of CCMV. CCMV solutions had 10 mg/L of K72 or GFP-K72 and aFT solutions 60 mg/L of K72 or GFPK72. The sample compositions are presented in Table S3 and obtained images in Figure S5. The samples were left to crystallize at 6 °C for 10 days before imaging. Samples with NaCl concentration of 100 mM or higher had no assemblies large enough to be detected by an optical microscope.
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Figure S5. Optical microscopy images of a) CCMV–K72, b) CCMV–GFP-K72, c)
aFT–K72 and d) aF-GFP-K72 in 40 mM NaCl solution.
Fluorescence Spectroscopy
The complex composition was studied by quenching the fluorescence of GFP-K72 via digestion of GFP using trypsin. Fluorescence of free GFP-K72 and that of CCMV and aFT complexes were measured for three hours in both absence and presence of trypsin. The measurements were conducted using an Agilent Technologies Cary Eclipse Fluorescence Spectrophotometer equipped with Varian PCB 1500 Water Peltier System. All the spectra were recorded in Brand semi-micro UV-Cuvettes (PMMA) at 37 °C using excitation wavelength of 400 nm. Sample compositions are presented in Table S4 and the measured spectra in Figure S6. The samples were prepared by combining all the components of each sample except for trypsin in a cuvette and mixing gently with a pipette. Trypsin was added last to the specified samples and solutions were stirred quickly with a pipette tip. All the samples were equilibrated in the sample holder for 5 minutes to reach the
133
measurement temperature before starting the measurement.
Table S4. Samples for the fluorescence spectroscopy
Figure S6. Fluorescence spectroscopy curves of GFP-K72, GFP-K72–CCMV and
GFP-K72–aFT complexes in both presence and absence of trypsin. The intensities are presented as percentage values of the initial intensity (t = 5 min after mixing) detected for each sample.
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Table S5. Advantage and Disadvantage of cocrystal assembly using SUPs
compared to synthetic polyelectrolytes.
Advantage Compared to conventional polyelectrolytes the number of charges and the charge density can be very well controlled in SUPs. Moreover, other proteins can be incorporated into the crystals by simple fusions. Compared to synthetic polylelctrolytes this would require an additional conjugation step.
Disadvantage Limited functionalities developed presently. In future, more customized functions might be studied with specific protein moieties. For instance, there are several examples representing (bio-)crystals that catalyze cascade reactions[48].
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