A Structure-Based Assembly Screen of Protein Cage Libraries in

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A Structure-Based Assembly Screen of Protein Cage Libraries in Living Cells: Experimentally Repacking a Protein-Protein Interface to Recover Cage Formation in an Assembly-Frustrated Mutant Thomas Cornell, Maziar S. Ardejani, Jing Fu, Stephanie Newland, Yu Zhang, and Brendan P. Orner Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.7b01000 • Publication Date (Web): 25 Dec 2017 Downloaded from http://pubs.acs.org on December 26, 2017

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Biochemistry

A Structure-Based Assembly Screen of Protein Cage Libraries in Living Cells: Experimentally Repacking a Protein-Protein Interface to Recover Cage Formation in an AssemblyFrustrated Mutant Thomas A. Cornell1,2, Maziar S. Ardejani1,2, † , Jing Fu2, ‡ , Stephanie H. Newland3, Yu Zhang2, § , Brendan P. Orner*1,2 1

Department of Chemistry, King’s College London, UK Division of Chemistry and Biological Chemistry, Nanyang Technological University, Singapore 3 School of Chemistry, University of Southampton, UK 2

Corresponding Author * [email protected] Current address: 15 Robin Road, Singapore

Present Addresses †

Scripps Research Institute, La Jolla, CA, USA Department of Biological Sciences, National University of Singapore, Singapore. § College of Chemical Engineering, Nanjing Forestry University, Nanjing, People’s Republic of China ‡

1 ACS Paragon Plus Environment

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ABBREVIATIONS

Dps, DNA Binding Protein from Starved Cells; OC/OB, one bead/one bacterium; FlAsH, Fluorescein Arsenical Helix(Hairpin) Binder; FACS, Fluorescence-Activated Cell Sorting; EDT, 1,2-Ethanedithiol; TPCR, transfer polymerase chain reaction; PI, propidium iodide; DIC, differential interference contrast microscopy.


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Biochemistry

ABSTRACT: Cage proteins, which assemble into often highly symmetric hollow nano-scale capsules, have great potential in applications as far reaching as drug delivery, hybrid nanomaterial engineering, and catalysis. In addition, they are promising model systems to understand how cellular nanostructures are constructed through protein-protein interactions, and they are beginning to be used as scaffolds for synthetic biology approaches. Recently, there has been renewed interest in the engineering of protein cages and, in support of these strategies, we have recently described a fluorescence-based assay for protein cage assembly that is specific for certain oligomerization states and symmetry-related protein-protein interfaces. In this current report, we expand this assay to living cells and a high throughput assay to screen protein cage libraries using flow cytometry. As a proof of principle, we apply this technique to the screening of libraries of a double alanine mutant of the mini-ferritin, DNA-binding protein from starved cells (Dps). This mutant, due to disruption of key protein-protein interactions, is unable to assemble into a cage. Randomization of residues surrounding the double mutation afforded a repacked interface and proteins with recovered cage formation, demonstrating the strength and utility of this approach.

KEYWORDS Ferritin, DNA binding protein from starved cells, protein-protein interactions, protein cage, protein library screening, FlAsH, protein engineering


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INTRODUCTION Members of the cage protein structural group assemble into hollow, nano-scaled, and often highly symmetric capsules. These include virus capsids1, vault proteins,2,3 and ferritins,4,5 among others.6 Due mostly to the presence of a central cavity which provides a sequestered environment and a solvent accessible surface distinct from the protein exterior, proteins in this class have a wide number of potential applications ranging from drug delivery7, catalysis8 and the synthesis of hybrid nanomaterials.9,10 In the latter direction, we have previously utilized ferritin protein cages to template the formation of extremely monodisperse preparations of gold nanoparticles,11 thus generating hybrid nanomaterials with the distinct advantages of increased water solubility, protection from Ostwald ripening, and the potential for straightforward engineering to present peptide affinity tags for directed delivery or for the construction of additional levels of nano-structural hierarchy. In the process of applying our approach to the further development of novel materials, it has become increasingly clear that these strategies can be limited by pernicious protein instability. Therefore we have strived to understand the fundamentals of the protein-protein interactions that control ferritin cage assembly12,13,14,15,16,17 and have discovered and characterized, through serendipity13 and through low-level computation-aided rational design,18,19 mutant ferritins with enhanced stability. While successful, these can be protracted strategies, and, thus, we thought that a diversity-based discovery approach to screen protein libraries could prove complimentary. It was envisioned that the most rapid and efficient high throughput screen of protein libraries would be one done with single library clones expressed in individual bacteria (we refer to these libraries as “one clone/one bacterium” (OC/OB) in analogy to “one bead/one compound” small molecule libraries20) with the bacteria screened using fluorescence-activated cell sorting (FACS) (Figures 1A and 1B).21 A limitation of traditional protein engineering methods employing library screening in living cells, is that they often couple the protein of interest to downstream enzymatic reporters or live/dead assays, sometime producing “hits” operating through undesired mechanisms. Because our interests are structural, and focus on one specific oligomerization state of a class of proteins that can form multiple states,16 we required a specific output that is directly tied to structure, and the use of FACS dictated that this output be fluorescence. The fluorescence-based nature of our recently developed cage-specific assay (described below) matches these requirements. Recently, we developed an assay, building on the work of the Tsien22 and Schepartz23 labs, that correlates fluorescence with protein cage formation.24,25 We have applied this assay to the E. coli mini-ferritin, Dps (“DNA-binding protein from starved cells”) and demonstrated that it works in lysates, can be scaled to medium throughput, and is specific for the cage oligomerization state in the presence of other states. The assay employs designed cysteine-rich, assembly-dependent sites across key protein-protein interfaces and the regent FlAsH which fluoresces upon binding to these sites. A first step to apply the assay to the screening of protein libraries (Figures 1A and 1B) was to establish that it could work in living cells and could be expanded to flow cytometry. This is one goal of the work described in this report. The second goal is described below.

Figure 1. (A) To establish protein cages with enhanced assembly ability, a high throughput screen of protein libraries was conducted. (B) Individual library members were expressed in single bacteria (“one clone/one bacterium” or OC/OB libraries) and screened for cage formation using our fluorescence, FlAsH-based assay24 coupled to cell sorting (FACS). (C) Designed variants used in this report depicted as names and schematics along with overall research strategy. (Panel C Top) Dps crystal structure (PDB: 1DPS) and all nanocage-forming designs and controls as described and characterized previously25: a positive control (DpsCCPGCC) with full tetracysteine FlAsH binding site, a design with a cage assembly-dependent FlAsH binding site (DpsPAGCC) and the negative control with no additional cysteines (Dps). (Panel C Middle) Mutation of both R83 and R133 to alanine results in Dps(AA)PAGCC, which has frustrated cage formation (see Figures 2A and 2B). (Panel C Bottom) FlAsH/FACS screening of an OC/OB library based on Dps(AA)PAGCC results in a mutant, Dps(AA)PAGCC*, with recovered assembly ability due to D130L, D142W, D143G mutations. In the protein schematics: cysteine (red), glycine (light grey), alanine (dark grey), proline (black).


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Biochemistry

The design of protein and peptide cages has been a recent interest of the protein engineering community,26,27, 28,29,30 and it potentially has future applications in the development of protein cages as synthetic biology scaffolds.31 Many of these design approaches are computation-based, and an experimental approach could prove complimentary.29 Therefore, as a second goal of this report, we set out to establish a proof of principle by determining whether it is possible to utilize our high throughput screen to effectively repack a protein-protein interface of a protein cage. Previously we have shown that the Dps double alanine mutation at positions R83 and R133, residues that are involved in an extended interfacial hydrogen bonding network near the three-fold axis of symmetry (Figures 2A and 2B), results in a mutant with disrupted cage forming ability and which cleanly assembles into only dimer. Therefore, we decided to apply our screen to a focused library based on this double alanine mutant. The randomized residues were positioned to surround the two alanines with the intention of discovering mutants with recovered cage-forming-ability by means of repacking the three-fold symmetric protein-protein interface. Not only would this objective, if successful, demonstrate a proof of principle of our methodology to generate protein cages with enhanced properties and assembly across specific symmetric interfaces, it would also provide insight into the fundamentals of the protein-protein interactions that control protein cage formation.

Figure 2. (A) In the E.coli Dps protein cage, R83 and R133 (red spacefill) are involved in a network of hydrogen bonds at a protein-protein interface near a three-fold axis of symmetry (B). Double mutation of these residues to alanine results in a mutant that has frustrated cage formation and cleanly forms a dimer.15 (C) Residues (N19, D20, V21, Q86, L87, N130, D141, D142, D143, S152, R153, and D156) (blue) surrounding the A83 and A133 mutations (the native arginine residues are highlighted in red) were randomized to generate libraries for the repacking of the interface to recover cage assembly. PDB: 1DPS.32 Note that Panel B depicts a different orientation from the others to aid visibility of the significant residues.

Materials and Methods Cloning, expression, purification, and characterization of proteins are described in detail in the supporting information. Live cell fluorescence microscopy Suspensions of the bacteria harboring expressed and labeled proteins (see Supporting Information) were resuspended (100 µL, FlAsH buffer, 4 °C). At this stage, dyes could be added to label all and nonviable bacteria (Hoechst 33342, 1 µL of a 5 mg/mL stock; propidium iodide, 2 µL of a 1 mg/mL stock; Invitrogen). The suspension (5 µL, 4 °C) was mounted on a polylysine slide with a coverslip and images were obtained on a wide field Nikon Eclipse Ti-E Inverted microscope at 100x magnification (Filter sets: Hoechst 33342: Ex - 402 ±15 nm, Em - 455 ±50 nm. FlAsH: Ex – 490 ±20 nm, Em – 525 ±36 nm. Propidium iodide: Ex – 555 ±25 nm, Em – 605 ±52 nm) and analyzed with ImageJ.33 Flow cytometric analysis Suspensions of the bacteria with expressed and labeled proteins (see Supporting Information) were resuspended. Additional dyes were added to label all and nonviable bacteria (Hoechst 33342, 1 µL of a 5 mg/mL stock; propidium iodide, 2 µL of a 1 mg/mL stock; Invitrogen). The suspension was filtered (BD) and injected onto a BD Fortessa (Hoechst 33342: Ex laser - 375 nm, Em filter - 450 ±20 nm. FlAsH: Ex laser – 488 nm, Em filter – 530 ±30 nm. Propidium iodide: Ex laser – 488 nm, Em filter – 670 ±20 nm) with data analyzed with FACSDiva and FlowJo (Tree Star) software. Hoechst positive cells were gated first to identify all bacteria. This population was further gated to identify the PI negative (i.e. living) cells, and these were analyzed for FlAsH fluorescence. OC/OB library generation through transfer PCR (TPCR)


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The TPCR34,35 library used Dps(AA)PAGCC in pET-46 as a template and included five forward primers with NNS codons at specific positions and along with one reverse primer (Table S3). The PCR reaction included Pfu reaction buffer (5 µL of 10X, Promega), dNTP mix (2 µL of a solution containing dATP, dTTP, dGTP, and dCTP at 2 mM each, Promega), forward and reverse primers (8 nM of F1 and F5, 6 nM of F2, F3, F4 and R1, IDT) dsDNA template (20 ng), DMSO (2% v/v), Pfu polymerase (1 µL of 3 U/µL, Promega) and deionised H2O (to a total of 50 µL), and this was subjected to an initial melting step (95 °C for 5 min) followed by 40 cycles of amplification (95 °C for 30 s, 58 °C (increased 0.3 °C every cycle) for 1 min, and 72°C for 6 min) followed by 72 °C for 10 min. The amplification solution (17 µL) was then treated with DpnI (1 µL of 10 U/µL, NEB) in a reaction containing NEB buffer 4 (2 µL of 10x) (37 °C, 1.5 h). The solution was desalted (PCR Cleanup Wizard, Promega) before transformation. The desalted constructs (8 µL) were electroporated (XL-1 Blue, Novagen) and the resulting transformed bacterial were recovered (SOC, 1 h, 37 °C), further incubated (37 °C, carbenicillin, 1 µL of a 50 mg/mL stock in 1 mL of SOC, overnight), and the plasmids were isolated via miniprep (Sigma). The desalted plasmids (PCR Cleanup Wizard, Promega) were electroporated into a protein expression host (8 µl (Rosetta, Novagen) recovered with SOC (1 ml, 1 h, 37 °C)). In total, fifteen transformations were performed and the resulting bacteria were pooled together and incubated (overnight, 37 °C, carbenicillin, 1 µl of a 50 mg/ml stock). Based on transformation efficiency, the overall diversity was estimated to be 63,000 clones. FlAsH/FACS screening of OC/BC libraries Pelleted (4,000 rpm, 2 min) library bacteria (see above) were resuspended (10 mL LB with 10 µL of a 50 mg/mL carbenicillin) and incubated (1 h, 37 °C), and protein expression was induced (IPTG, 5 µL of a 1 M stock, 30 °C, 1.5 h). For each round of screening, cultures were prepared and labeled as above for the flow cyotmetric analysis and injected (for the first round, a total of 4 mL in 1 mL batches) onto a BD Aria III and sorted (Hoechst 33342: Ex laser - 375 nm, Em filter - 450 ±20 nm. FlAsH: Ex laser – 488 nm, Em filter – 530 ±30 nm. propidium iodide: Ex laser – 488 nm, Em filter – 670 ±20 nm) with data analyzed by FACSDiva and FlowJo (Tree Star) software. Hoechst positive cells were gated first to identify all bacteria. This population was further gated to identify the PI negative (i.e. living) cells, and these were further analyzed for FlAsH fluorescence with the highest intensity cells collected. This later population was gated so that only 1% of the total number of live cells was collected. The collected cells were suspended (10 mL total of LB with 10 µL of a 50 mg/mL carbenicillin stock) and incubated (overnight, 37 °C). After the final sort, the suspension was plated on LB agar plates (50 µL/mL carbenicillin, 34 µL/mL chloramphenicol) and selected colonies were submitted for sequencing. RESULTS Protein Design and In Vitro Characterization For the optimization of the FlAsH-based assay in live cells, previously characterized controls were used (Figure 1C). These included the cage forming protein DpsCCPGCC, which has a tetra-cysteine motif on every monomer and thus generates a strong FlAsH signal that is not oligomerization dependent. We also used the wild-type protein Dps which can form a cage but generates low fluorescent signal because it has no FlAsH binding site. Our further designs were all based on DpsPAGCC which presents two cysteines at the monomer C-terminus and forms a cysteine rich, FlAsH binding site only upon cage formation. Based on this protein, we also designed Dps(AA)PAGCC, which contained the double mutation R83A/R133A. Our previous work15 (see above) showed that this mutant would not assemble into a cage because of a disrupted network of salt bridges (Figures 2B and 5A) and, because it presents a cage-dependent FlAsH binding site, should not bind FlAsH. As further controls, the double tryptophan and double aspartic acid mutants were also generated at the same positions (see Supporting Information). The double alanine protein, Dps(AA)PAGCC, was used as a basis for library design (see below). All proteins (Figure 1C), including the library (see below), were expressed from a pET-46 Ek/LIC plasmid which provides a small N-terminal His6 affinity tag separated from the protein by a short enterokinase protease cleavage sequence. This strategy permitted in vivo selection and in vitro characterization of purified proteins without the need for recloning.24 Upon expression and purification (Figure S1, Table S5), the cage forming designs, DpsCCPGCC, DpsPAGCC, and Dps, all were determined to form the cage state predominately, and the cage-frustrated designs, including Dps(AA)PAGCC, formed smaller particles in dynamic light scattering (DLS), did not form cage as determined by transmission electron microscopy (TEM), and only assembled into dimers as evidenced by size exclusion chromatography (SEC) (Figures S2, S3, S4, S14, 4A, and 4B) These results were consistent with the trends observed in the signal generated through FlAsH binding (Figure S5). Of the double mutants at positions 83 and 133, Dps(AA)PAGCC generated the lowest signal, and therefore this mutant was carried forward as the basis for library generation. Optimization of Cage Assembly Detection with FlAsH in Living Cells The optimization of conditions for applying our FlAsH-based cage assay to living cells was modified from Nhieu and coworkers36,37 and Gierasch and co-workers38 and primarily focused on the adjustment of FlAsH concentration to minimize toxicity and maximize signal, addition of redox active reagents to maximize signal and minimize background, and incubation times to maximize signal and reproducibility. Epi-fluorescence microscopy resulted in images that correlated with the trends observed in the in vitro work (see above). Bacteria expressing the protein with an oligomerization-dependent FlAsH binding site, DpsPAGCC, generate fluorescence that is intermediate between that of cells expressing the positive and negative control proteins, DpsCCPGCC and Dps, respectively (Figures 3A, S6, and S7). These conditions were applied to, and further optimized for, flow cytometry. The dyes, Hoechst 33342 and Propidium iodide (PI), were used to first gate for all bacteria (Hoescht 33342 positive cells) and living bacteria (PI negative cells) before FlAsH analysis. Propidium iodide stains DNA but cannot pass through the membrane. Therefore cells with ruptured membrane (i.e. dead


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Biochemistry

cells) will be PI positive. Consistent with all the previous in vitro and in vivo controls, the population of bacteria expressing the protein with the oligomerization-dependent FlAsH binding site, DpsPAGCC, generates an average fluorescence that is intermediate between that of the populations of cells expressing the positive and negative controls, DpsCCPGCC and Dps, respectively (Figure 3B). Furthermore, the population of bacteria expressing the cage-frustrated mutant, Dps(AA)PAGCC, exhibited a relatively weak fluorescence that was an order of magnitude lower than that of DpsPAGCC (Figure 3C), strongly demonstrating the feasibility of using flow cytometric analysis to screen libraries based on the double mutant. Building upon these robust controls and initial procedures, we set out to establish the screening technique.

Figure 3. FlAsH-based assay in living cells to determine degree of cage assembly. (A) Determination of intracellular FlAsH labeling of overexpressed Dps protein controls in E. coli with epi-fluorescence microscopy comparing positive and negative controls, DpsCCPGCC and Dps respectively, with a protein presenting a cage-dependent binding site, DpsPAGCC. Note: Differential interference contrast (DIC) micrographs are provided to image both fluorescent and non-fluorescent bacteria. DIC image: 100x magnification, 20 ms exposure. Green channel: 50 ms exposure, filter sets: Ex 490 ±20 nm, Em 525 ±36 nm. (B) Determination of the oligomerization state of Dps controls using flow cytometry. Distribution of fluorescence signal for populations of cells expressing negative control, Dps (light grey), positive control, DpsCCPGCC (black), and protein displaying cage-dependent FlAsH binding site, DpsPAGCC (red). (C) Determination of the oligomerization state of the cage-frustrated design, Dps(AA)PAGCC (blue), compared to its parent protein which does form cage DpsPAGCC (red), using flow cytometry. Populations were gated for Hoechst 33342+ and PI-. Blue channel: Ex laser 375 nm, Em filter 450 ±20 nm. Green channel: Ex laser 488 nm, Em filter 670 ±20 nm. Data was normalized (FlowJo, Tree Star) using 100,000 cells. Schematics of proteins match those defined in Figure 1C.

Library Design, Generation, Screening, and Hit Characterization As our goals were primarily structural and specifically focused on protein-protein interfaces, we took a directed approach to library generation. Instead of the more common error prone PCR39 which is widely used for enzyme evolution40 and generally produces mutations throughout a gene, we employed transfer PCR (TPCR).34 This method allows the targeting of specific sites in the gene so that key residues can be combinatorially mutated. Moreover, directed mutation permits the generation of libraries that have a large population of clones with multiple mutations. This aspect is important for the goal of interface repacking because residues involved in protein-protein interactions are thought to be highly cooperative.41 To determine the positions to randomize during library generation, a contact analysis of residues near R83 and R133 in the crystal structure (1DPS32) of the wild-type protein was performed (Figure S8) and the nine residues, N19, D20, V21, D141, D142, D143, S152, R153 and D156, were identified as forming a “contact network”.42 These amino acids reside at three distinct positions in the tertiary structure of the Dps monomer four-helix-bundle (defined by helices A-D). The first set of residues, N19, D20 and V21, are located on the N-terminal tail leading to the A-helix. The second set of residues, D141, D142 and D143 are at the Nterminus of the D-helix. Third, S152, R153 and D156 are also on the D-helix but across the interface from D141, D142, and D143 of another monomer.


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Further inspection of the Dps crystal structure showed that Q86, L87 and N130, while not involved directly in the contact network, are positioned spatially proximal to the network, making them additional candidates for randomization. The residues Q86 and L87 are at the end of the B-helix C-terminus and N130 is positioned along the C-helix. Taken together, these represent a large proportion of the residues located at the protein-protein interface, and all were randomized, while maintaining the two alanine mutations at positions 83 and 133, for the generation of the library based on Dps(AA)PAGCC (Figures S9 and 2C) After four rounds of FlAsH/FACS screening, twenty clones were sequenced (Table S4) and three were taken forward for purification and characterization (Figure S10, Table S5) based on either their similarity to other sequenced clones or on their uniqueness. A single mutant (N130Y) was selected because a mutation at this position was observed for the majority of the clones, and this specific sequence was highly represented in the matured library. A triple mutant (Q86V, L87F, N130F) was taken forward because hydrophobic and aromatic residues tend to be energetically important residues in protein-protein interactions. Both these mutants expressed poorly and proved difficult to purify (Figure S10). A third mutant was selected (N130L, D142W, D143G which also contained a silent mutation at position 141) because it was the only sequenced clone with mutations at positions 142 and 143. This protein expressed robustly and could be purified to near homogeneity, and it became our primary focus. It will be referred to hereafter as Dps(AA)PAGCC*. Extensive characterization of the purified Dps(AA)PAGCC* demonstrates that it has recovered cage assembling ability. Assessment of cage formation using our in vitro FlAsH assay indicates a significant increase in fluorescence signal over the crippled parent protein, Dps(AA)PAGCC (Figure S11). Furthermore, DLS indicates that Dps(AA)PAGCC* forms particles that are significantly larger than the frustrated mutants such as Dps(AA)PAGCC and on par with those formed by the cage forming proteins, DpsPAGCC, DpsCCPGCC, and Dps (Figures 4A and S14). In addition, SEC suggests the presence of a significant population of the cage state in Dps(AA)PAGCC* compared to undetectable amounts in Dps(AA)PAGCC. (Figures 4B, 4D, and S12, and S13) Interestingly, SEC analysis of replicate preparations of Dps(AA)PAGCC* (Figures 4D and S13), while still exhibiting a significant population of the cage state, suggests that the ratio of assembly is more variable than related proteins with which we have worked. In addition, along with cage and dimer states, an additional assembly state is observed. Its retention volume is consistent with a complex that is intermediate in size between a dimer and the fully assembled cage. Finally, negatively stained TEM indicates clear cage structures with diameters consistent with the wild-type protein (Figures 4C and S15). It should be noted that this trend of cage recovery is consistent with that seen in the characterization of the other two expressed clones (N130Y and Q86V,L87F,N130F) obtained from the library screen (Figures S11, S12, S14, and S15), but, purification challenges (Figure S10) make it difficult to reach similar conclusions with comparable confidence. Taken together, these data strongly demonstrate that Dps(AA)PAGCC* forms a substantial population of protein cage and has recovered this ability through a repacked protein-protein interface discovered from a FlAsH/FACS screen of the OC/OB library.

Figure 4. A mutant with a repacked protein-protein interface discovered from a FlAsH/FACS screen of the OC/OB library, Dps(AA)PAGCC*, has recovered cage assembly properties. (A) Dynamic light scattering of Dps(AA)PAGCC* compared to cageforming and cage-frustrated controls. Error bars indicate S.E. Note: these values are solvated hydrodynamic diameters. (B) Size exclusion chromatography of cage-forming and cage-frustrated controls, DpsPAGCC and Dps(AA)PAGCC, respectively. (C) Transmission electron micrograph of cage-recovered mutant Dps(AA)PAGCC*. Note: this sample is negatively stained and under vacuum. Scale bar indicates 20 nm. (D) Peak distributions of Dps(AA)PAGCC* for assembly states identified by size exclusion chromatography. Average of five protein preparations (Figure S13). Error bars indicate S.E. Schematics of proteins match those defined in Figure 1C.

DISCUSSION Although our extensive characterization shows that Dps(AA)PAGCC* is forming cage, and thus demonstrating that it is indeed possible to repack a protein-protein interface using our technique, it is worth highlighting some subtle SEC observations. From five protein preparations, we see quite a wide distribution of ratios between the dimer and cage states for the protein (Figures 4D and S13). Furthermore, we observe an additional intermediate that is larger than a dimer but smaller than the cage. Although a


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Biochemistry

full analysis of the kinetics of assembly of Dps(AA)PAGCC* is beyond the scope of this paper, this variability could be the result of enhanced assembly dynamics or population exchange. Moreover, it is notable that the residues chosen for randomization in the library primarily reside at the three-fold, trimerization interface of the cage. It is reasonable that these modifications would enhance this interface’s stability or lifetime during assembly. This enhancement could be simply energetic or an amplification of a subtle structural alteration akin to the “arch and keystone” effect which we originally put forth with respect to a dimer intermediate.18 More generally, this effect relates to structural modification at a protein-protein interface that further influences the relative projections of distal protein surfaces.26,43 The alteration of these surfaces with respect to each other affects the assembly of closed structures much like how changing the relative angles of an architectural keystone influences the structure of an arch built around it. Thus, this “arch and keystone” effect could be playing a role in the formation or further assembly of the Dps(AA)PAGCC* intermediate, extending its lifetime. Consequently, further investigation into the mechanism of assembly of this mutant could shed light on the various energetically accessible pathways to assembly. Modeling44,45,46 of the re-packed three-fold symmetric, trimeric interface of the Dps(AA)PAGCC* protein cage (Figure 5) provides possible insight into the protein’s recovered assembly and the role the mutations play in it. While we employ widely adopted computational techniques, it should be noted that the resulting calculated structure is not a high-resolution crystal structure and should be treated as a theory to explain the fact the mutant pulled from the cage-specific screen has had its assembly irrefutably recovered through only three mutations. We hope that this model can help direct next steps in further research. The mutated residues in the model not only are predicted to make new interactions compared to Dps (Figure 5A), but they rearrange the interaction network across the interface. Along with R83 and R133 which were mutated to alanine in the parent of the library, Dps(AA)PAGCC, a number of native residues moved out of (V21, Y16, S152 and I166) and some new ones were brought into (N19, A145 and D146) the network of interactions. Although it is well-accepted that a protein-protein interface is highly cooperative and its stability is due to more than the sum of its parts41, some insight, in this case, can be gleaned from inspection of reduced sets of interactions. Most striking are the edge to face interactions47,48 predicted to be made by the D142W where three tryptophan indole side chains are modeled to pack against each other at the three-fold axis of symmetry (Figure 5B). It is tempting to draw parallels to a previously published design of the maxiferritin e. coli bacterioferritin (Bfr).18 In that study, we found that by replacing an aspartic acid residue with the aromatic amino acids (tryptophan, tyrosine, or phenylalanine were equally successful), the cage/dimer mixture could be pushed toward 100% cage. Modeling suggested a symmetric edge to face interaction, similar to that proposed in our calculated structure (Figure 5A), between the three aromatic residues across a three-fold axis near the monomer N-termini. A significant difference between these studies was that the current design relies on an experimental combinatorial approach whereas the Bfr design was mainly computational. Additionally, the Bfr cage is a 24-mer with octahedral symmetry, with a single set of equivalent three-fold trimeric interfaces, whereas Dps is a tetrahedral 12-mer with two distinct sets of three-fold interfaces--one centered around the monomer C-termini (the “Dps-like” interfaces) and the other around the N-termini (the “ferritin-like” interfaces). The D142W mutations in Dps(AA)PAGCC* are located at the N-terminal, ferritin-like interface. Moreover, although tertiary and quaternary structure of Dps and Bfr are poorly superimposable, the spatial projection of D142W and the aromatic residues in the stabilized Bfr designs are reminiscent. Comparing these two successful studies might suggest broad design rules for the formation of general three-fold symmetric homotrimeric interfaces, or the potential of shared structural energetics for a class of structurally divergent cage proteins with related cellular functions (i.e. the ferritins). However, this report is intended as an initial proof of principle demonstrating a new experimental technique to discover protein cage mutants with modified structural properties. Thus, it should be pointed out that only a small percentage of theoretical diversity was explored by our library, the library generation has not been fully optimized, and the structural explanations we are providing are based on modeling. With that said, these speculations could provide a direction for future inquiry. The other two mutations in Dps(AA)PAGCC* can be understood with respect to D142W. For the symmetric edge to face interaction of the tryptophans to be achieved, an interfacial rearrangement is necessary. Modeling suggests that this rearrangement is sterically facilitated by the D143G and N130L mutations which are proposed to be involved in changes with interactions to the polypeptide backbone. With respect to wild-type, the calculations show that the backbone interactions with the N130 residue are broken and new backbone interactions are formed with G143 (Figures 5C and S16). It is useful to place the structural consequences of these mutations into context by noting conceptual differences between strict rational design and our directed combinatorial approach. The three-fold edge to face interaction made by D142W is the type that can be recognized clearly and thus relatively easily rationally designed.18 However, rearrangements and complex conformational changes that facilitate this interaction would be much more difficult to engineer and it is unlikely that D143G and N130L could have been found through a purely rational strategy. This is especially true for alterations of the backbone. Thus, the ability to discover unexpected and serendipitous packing solutions emphasizes the power of experimental combinatorial approaches. CONCLUSIONS The research presented in this report carries our assembly assay for specific protein cage oligomerization states into living cells and couples this technology to the screening of high throughput libraries of protein cages. The strength of this assay is that it is structurally dependent, and we’ve demonstrated that this technique is specific not only for assembly but for a specific assembly state. Through the research presented herein we provide strong support that assembly in protein cages can be recovered, that protein-protein interactions in large assemblies can be experimentally repacked, and that our screening strategy can select for specific properties of protein cages from randomized libraries. We think this approach will prove complimentary to computational ap-


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proaches to design proteins cages30 and will help shed light on the fundamentals of protein-protein interactions thus facilitating future protein engineering strategies.

Figure 5. The predicted interface of Dps(AA)PAGCC* that was repacked by mutations (D130L, D142W, D143G) discovered through library screening. (A) Calculated interaction maps of two monomers of both DPS and Dps(AA)PAGCC*. Grey residues are mutations causing disruption of cage formation and green residues indicate mutations established from the library screen; bold circles indicate residues that were removed or brought into the interaction network upon mutation. Black, green, red, and blue lines indicate van Der Waals, ionic, aromatic, and hydrogen bond interactions, respectively, and an arrow indicates sidechain-to-backbone interactions. Thicker lines indicate interactions that are not present in wild-type and are a consequence of mutation and re-packing. To reduce complexity, interactions with residues in the same or with a third monomer were omitted. (B) Predicted interaction with the three-fold symmetric D142W edge-toface interaction and (C) further predicted interactions near D142W and D143G. The predicted structure was based on 1DPS32 and calculated with FoldX44 and Rosetta45. The residue interaction map was generated with Aquaprot46. Schematics of proteins match those defined in Figure 1C.

ASSOCIATED CONTENT Supporting Information In a single PDF file, details for the cloning, expression, purification, and characterization of proteins are described in detail along with details for preparation of bacteria for fluorescence microscopy and flow cytometry, sequences for primers, SDS-PAGE gels, size exclusion chromatograms. transmission electromicrograms, and fluorescence values after FlAsH labeling for purified proteins. Additional fluorescence micrographs are provided along with contact analysis of the DPS interface and additional images of positions that were mutated. Sequences of clones isolated from the screens and characterization (SDS-PAGE, MALDI-TOF, fluorescence after FlAsH labeling, SEC, DLS, and TEM) of the purified proteins are provided along with additional modeling figures of Dps(AA)PAGCC*. Full sequencing results for all genes used in this study are also provided.

AUTHOR INFORMATION Author Contributions The manuscript was written through contributions of all authors. / All authors have given approval to the final version of the manuscript.

Funding Sources T.A.C was sponsored by SINGA and BSE scholarships at NTU and King’s respectively. The research at NTU was supported by an SPMS start-up grant, a Singapore Ministry of Education Academic Research Fund Tier 1 Grant (RG 53/06) and B.P.O’s personal salary. At KCL, it was supported by a Marie Curie CIG: PCIG13-GA-2013-618538 and B.P.O’s personal salary.

ACKNOWLEDGMENT We thank M. Peczuh, N. Luedtke, and Fan Rongli for insightful conversations. We also thank D. Thurston’s lab for help and access to instrumentation, as well as the technical staff in both KCL’s Centre for Ultrastructural Imaging and the Guy’s and St Thomas’s flow cytometry centre for help in TEM and flow respectively. We also thank Bobbi Fleiss for her help in FACS analysis.


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Biochemistry

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