Controllable Assembly of Flexible Protein Nanotubes for Loading

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Biological and Medical Applications of Materials and Interfaces

Controllable Assembly of Flexible Protein Nanotubes for Loading Multifunctional Modules Guibo Rao, Yan Fu, Na Li, Jiayi Yin, Jie Zhang, Manli Wang, Zhihong Hu, and Sheng Cao ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b07611 • Publication Date (Web): 10 Jul 2018 Downloaded from http://pubs.acs.org on July 12, 2018

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ACS Applied Materials & Interfaces

Controllable Assembly of Flexible Protein Nanotubes for Loading Multifunctional Modules ‡

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†

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Guibo Rao,†, Yan Fu, Na Li, ,‡ Jiayi Yin, ,‡ Jie Zhang, ,‡ Manli Wang,§ Zhihong Hu,§ Sheng Cao

†

*

†

CAS Key Laboratory of Special Pathogens and Biosafety, Center for Emerging Infectious

Diseases, Wuhan Institute of Virology, Chinese Academy of Sciences, Wuhan 430071, PR China ‡

University of Chinese Academy of Sciences, Beijing 100049, PR China

§

State Key Laboratory of Virology and Joint-Lab of Invertebrate Virology, Wuhan Institute of

Virology, Chinese Academy of Sciences, Wuhan 430071, PR China *

Correspondence to: [email protected]

KEYWORDS: nanotube, self-assembly, helical reconstruction, cryo-EM, functionality

ABSTRACT

Viruses with filamentous morphologies, such as tobacco mosaic virus (TMV) and M13 bacteriophage, have long been studied as multivalent nano-scaffolds for loading functional

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motifs. Structural assembly of the capsid proteins (CPs) of filamentous viruses often requires the presence of DNA or RNA molecules, which has limited their applications. Here, we describe a strategy for controllable assembly of flexible bio-nanotubes consisting of Escherichia coli expressed CP of baculovirus Helicoverpa armigera nucleopolyhedrovirus (HearNPV) in vitro. These protein-only nanotubes were studied as a new structural platform for high-density presentation of multiple active molecules on the exterior surface by direct fusion of the protein of interest to the N-terminus of HearNPV CP (HaCP). Structural characterization using cryoelectron microscopy (cryo-EM) demonstrated that HaCP could assemble into two closely related but structurally distinct tube types, suggesting the tunable HaCP interaction network is the major contributor to the flexibility of HaCP nanotubes. Our flexible nanotubes could tolerate larger molecular modifications compared with TMV-based templates and could be used as promising candidates for versatile molecular loading applications.

INTRODUCTION Viruses are natural masters of programmed self-assembly, and inspired by these natural selfassembly systems, various artificial supramolecular complexes have been developed in materials design and synthesis in recent decades.1-3 Multiple copies of capsid protein (CP) self-assemble into viral capsids in a highly-ordered fashion, thus offering attractive building blocks for the immobilization and presentation of functional modules at nanoscale resolution. Most studied platforms in CP-based nanotechnology are plant or bacterial viruses organized in either icosahedral or helical symmetries, appearing as spherical or filamentous morphologies, respectively. Compared with icosahedral capsids, which usually have hundreds of modifiable


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CPs, helical assemblies can provide thousands of identical CPs as templates for genetic or chemical modifications, thus serving as more powerful multivalent nano-templates. Filamentous viruses can be generally classified as flexible viruses, such as M13 bacteriophage, and rigid rodshaped viruses, such as tobacco mosaic virus (TMV). They exhibit extended exterior surfaces and have been used as nano-scaffolds for displaying engineered peptides.4-5 A direct way to harvest functionalized nanoparticles assembled in host cells is to extend or insert the coding sequence of a protein of interest into viral CPs by genetic manipulations. This one-step incorporation approach may appear convenient and straightforward, however genetic modifications of CPs often disturb or even abolish assembly of the ordered bio-scaffolds due to steric hindrance between bulky CP fusion proteins.3 To form a normally assembled virus particle, TMV CP (158 aa in length, ~2130 copies per virion) can tolerate an extension of less than 20 amino acids at the C-terminus,6 and pVIII of M13 (50 aa, ~2700 copies) can carry an N-terminal extension of up to 10 residues.7 Two general strategies have been applied to increase the loading capacity of helical scaffolds. One approach is to connect functional moieties to genetically or biochemically tailored amino acid side chains exposed on pre-formed capsids through a conjugation reaction.8-9 An alternative strategy is to produce partly decorated virus particles by reducing the copy number of the protein of interest on the capsid through genetic manipulations.4, 10

The reproductive power of viruses has made these virus-based systems a potential economic

choice for many applications; however, variations of the viral surface may also induce necrotic responses in the host and lower the yield of virus products,11 which would limit virus-based applications. To develop more applicable bio-templates, viral CPs have been studied as building blocks for helical assembly in vitro.12-13 TMV has been the most studied model system for virus assembly


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in the presence of an RNA molecule containing a so-called “origin of assembly sequence” (OAs).14 Multiple TMV CP mutants can co-assemble with RNA into rigid nanotubes, which may serve as promising scaffolds for loading various functional motifs simultaneously.15 However, addition of genomic materials would not only complicate the preparation procedure, but also limit applications owing to safety concerns. Although purified TMV CP alone could assemble into nanotubes at pH 5.5,16 the applications are highly limited owing to loss of function for many proteins in acidic conditions. Therefore, new helical materials, which could be controllably assembled from a CP component alone, under mild physiological conditions, would be highly desirable. The helical biomaterials we describe here are nanotubes assembled from the recombinantly expressed CP of Helicoverpa armigera nucleopolyhedrovirus (HearNPV). HearNPV, a group II Alphabaculovirus in the family Baculoviradae, has been successfully used as viral pesticides for controlling cotton bollworm infestations.17 Baculoviruses are rod-shaped particles, which gave rise to their family name: Baculoviridae (the Latin term "baculum" means stick or staff).18 Baculovirus capsid is made of the major CP and a number of other minor capsid-associated proteins.19 The major CP is one of the most abundant proteins in mature virions,20 and is conserved in all baculovirus genomes

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(Figure S1A). Interestingly, aberrant long electron-

lucent nanotubes were found in the host cells when a variety of viral genes had been knocked out.22-28 The major CP was the only detectable component of the tubes, into which no genomic materials were correctly packaged.27 We set out to develop a new tubular bio-template based on these unique self-assembled bio-nanotubes. For simplicity, the major CP of HearNPV will be hereafter referred to as HaCP.


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RESULTS AND DISCUSSION In vitro assembly of HaCP nanotubes In order to achieve HaCP self-assembly into nanotubes, we developed a three-step strategy that involves the controllable activation of assembly-incompetent HaCP into assembly-competent states (Figure 1A). First, to block the uncontrollable self-assembly process, HaCP was genetically modified with an N-terminally fused protein tag and was expressed in Escherichia coli. We found that the GST tag provides sufficient steric hindrance to block self-assembly of HaCP in the cells, and the soluble fusion protein could be harvested from the cell lyse. Purified GST-HaCP oligomerized into soluble light aggregates, not tube-like structures (Figure S2A). In the second step, GST-HaCP in the assembly buffer (see MATERIALS AND METHODS), bound to glutathione agarose resin specifically, as the standard affinity purification protocol suggested. Following the binding step, assembly-competent HaCP was released from the affinity resin using 3C protease, targeting the linker region between GST and HaCP. Once HaCP had been cleaved from GST, the HaCP nanotube could self-assemble readily in the assembly buffer at 4 °C overnight (Figure 1A).


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Figure 1. Assembly of HaCP nanotubes in vitro. A, B) Schematic flowchart for preparation of HaCP nanotubes (A) and EGFP-functionalized nanotubes (B). There are three types of recombinant protein expressed in E. Coli cells: GST-HaCP, GST-3C, and GST-EGFP-HaCP. During the three-step assembly of HaCP nanotubes, HaCPs are in three forms: free GST-HaCP, resin-bound GST-HaCP, and flexible nanotubes consisting of HaCP. To obtain functionalized nanotubes, free GST-HaCP and GST-EGFP-HaCP are mixed before being applied to the glutathione resin. C) TEM image of negatively stained HaCP assemblies showing overall flexible tubular structure. Scale bar: 500 nm. D) Heavily stained HaCP nanotubes at 50,000× magnification. White arrows indicate the characteristic banding densities in the nanotube. Scale


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bar: 50 nm. E) Lightly stained HaCP nanotubes at 29,000× magnification. Arrows indicate righthanded helical lines on the tube surface. Scale bar: 100 nm.

Size exclusion chromatography (SEC) with a pre-packed GE Sephacryl S-500 HR column was performed to purify HaCP nanotubes (Figure S3A–B). HaCP nanotubes passed through the packed bed and were eluted first following the void volume using PBS as the mobile phase. The yield of purified HaCP nanotubes was ~1 mg per liter of bacterial culture. Other components, identified as GroEL from E. coli (Figure S3C–D) and irregular HaCP oligomers, were eluted later in order of decreasing size. GroEL is a major molecular chaperon responsible for the correct folding of ~10% of the bacterial proteome.29 It is likely that GroEL was able to interact with partially folded GST-HaCP and the complex could be enriched by glutathione agarose. Purified HaCP nanotubes were negatively stained with 2% phosphotungstic acid (PTA) and imaged using TEM. When the stain had diffused into the nanotubes during an extended staining process (300 s), the tubes appeared as bent cylindrical shells surrounding a hollow channel (Figure 1C). 80% of the tubes ranged from 1 to 4 µm in length (Figure S2D–E). At higher magnifications, the wall densities appeared varied in thickness (Figure 1D), resulting largely from drying artifacts during negative staining. Close examination of the tube densities revealed that the banding patterns between sidewall densities run along the long axis of the tube with almost constant intervals (Figure 1D, inset). In contrast, when only a small amount of stain was absorbed onto the tube, the nanotubes appeared as lightly stained long cylinders with righthanded helical lines on the surface (Figure 1E, inset). Helical arrangement within the nanotube


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assemblies was thus evidenced by these image patterns, which probably arose from the penetration of stain into a set of grooves on the interior or exterior surface of the nanotubes. Unlike for TMV nanotubes, nucleotide sequences were not required for the assembly of HaCP nanotubes, and introduction of DNase I into the assembly system to remove DNA contamination had no apparent impact on the morphologies of the nanotubes (Figure S2B). Previous studies suggest that baculovirus DNA is bound to and condensed by dephosphorylated P6.9 during viral DNA encapsulation.28 When the heterogeneous mixture of purified bacmid DNA and P6.9 was introduced into the assembly system, the nanotubes also formed normally (Figure S2C). In the absence of nucleotide sequences, TMV CPs were assembled as two-layer disks at neutral pH, and acidic pH triggered the transition of disks into helices.30 In contrast, HaCP nanotubes without DNA largely maintained structural integrity at pH values ranging from 4 to 9 (Figure S4). Stronger acidic (pH9) treatment of HaCP nanotubes depolymerized the tubular arrangement, and this depolymerization process was irreversible when pH was adjusted back to neutral. Taken together these findings suggest that HaCP-only nanotubes could serve as an attractive molecular carrier in physiological or mildly acidic microenvironments.

Multi-Functionalized HaCP nanotubes Fluorescent proteins (FPs), widely used as imaging labels in life science, have also been used as standard modules to test the loading capacity of various CP-based platforms, such as M13,8 Potato virus X,31 and TMV.9 Four FPs were used in this study: mTagBFP (blue, excitation 399 nm/emission 456 nm), EGFP (green, 488 nm/509 nm), DsRed (red, 556 nm/586 nm), and mKate (far-red, 588 nm/635 nm) (Figure S1B). Fluorescent labels have been appended to the N- or C-


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terminus of baculovirus CPs for visualizing virus movement into host cells in vivo.32 However, we found that when FPs were fused to the C-terminus of GST-HaCP, the fusion proteins were expressed mostly as insoluble inclusion bodies in E. coli. When FPs were inserted between GST and HaCP, higher soluble protein yield was achieved. After the mixture of GST-FP-HaCP and GST-HaCP was applied to affinity resin, the resin-bound proteins were digested by 3C protease simultaneously (Figure 1B). FP-HaCP/HaCP hybrid nanotubes were then assembled similarly to the HaCP-only nanotubes. EGFP-HaCP/HaCP nanotubes were probed with anti-EGFP polyclonal antibodies and goldconjugated secondary antibodies. Immunogold-labeled HaCP nanotubes could be visualized using TEM (Figure 2A), suggesting that accessible EGFP molecules were distributed on the exterior surface of the tubes, since antibodies and gold particles (~10 nm in diameter) are too large to penetrate the wall of the nanotubes. To visualize EGFP-labeled nanotubes directly, fluorescent nanotubes were observed using a confocal microscope at different time points. Fluorescent signals were clearly enriched on the nanotubes and were evenly distributed along the entire length (Figure 2B). The fluorescent emission of EGFP-HaCP/HaCP nanotubes remained unchanged after a 5-day incubation at 4 °C (Figure 2C), demonstrating the high stability of the functionalized nanotubes. When HaCP, EGFP-HaCP, mTagBFP-HaCP, and DsRed-HaCP (at a molar ratio of 6:1:1:1, respectively) were co-assembled using the same mixing strategy, hybrid HaCP nanotubes were also readily formed. Co-localization of the three FPs on the tri-colored nanotubes was confirmed by confocal microscopy (Figure 2D), demonstrating that a combination of multiple functional moieties could be introduced at the point when the nanotubes were assembled.


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Figure 2. Incorporation of different proteins into the HaCP nanotubes. A) EGFP on the nanotubes was located with anti-EGFP antibodies and 10 nm colloidal gold-conjugated secondary antibodies. Scale bar: 1 µm. B) Confocal micrograph of EGFP-HaCP/HaCP nanotubes after 5-day incubation. Scale bar: 30 µm. C) EGFP emission strength was retained after 120 h, showing the stability of the functional groups on the hybrid nanotubes. N=5. NS, no significant difference (p>0.05). D) Confocal micrographs in three channels show the FPs, red DsRed, green EGFP, and blue mTagBFP, were introduced with high efficiency into the HaCP nanotubes simultaneously. Merged image demonstrates the co-localization of the FPs. Scale bar: 10 µm. E)


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Western blotting showing the different incorporation ratios for mKate-HaCP/HaCP hybrid nanotubes (mixing ratio of mKate-HaCP to HaCP was 1:2 (Lane 1), 1:4 (Lane 2), 1:8 (Lane 3), and 1:16 (Lane 4)). Anti-HaCP antibodies were used to identify and quantify mKate-HaCP and HaCP components. F) Western blotting of purified nanotubes. Lane 1: EGFP-HaCP/HaCP, Lane 2: nanoKAZ-HaCP/HaCP, Lane 3: HaCP only. Anti-HaCP antibodies were used to identify and quantify HaCP components. G) Luminescence patterns of the oxidation of coelenterazine with immobilized or free nanoKAZ. 0.005 nmol of nanoKAZ immobilized on HaCP were stable (for 5 days) and had higher catalytic activity than 0.005 nmol of free nanoKAZ.

For many of the perceived applications, it would be important to precisely control the incorporation rate of the desired functionalities.33 Different mixing/input molar ratios of mKateHaCP to HaCP were examined for this purpose. Western blotting analysis showed that mKateHaCP incorporation/output ratio decreased as the input ratio decreased (Figure 2E, and Figure S5A), suggesting that the incorporation could be performed in a predictable way. The output ratio of mKate-HaCP to HaCP in the hybrid tubes was estimated to be approximately 1:4 when the input ratio was 1:2 (Figure 2E, Lane 1). Although higher input ratios were tested, no more mKate-HaCP could be loaded (data not shown). Similar output ratios were found for other FPs (Figure 2F, Lane 1). Local defects on the functionalized nanotubes caused by unfavorable conformational changes or insufficient structural freedom on tagged HaCP molecules, could require unmodified HaCP to buffer the defects. We reasoned that tagged HaCP molecules could spatially separate from each other and thus reduce the steric interference between them, and the requirement for separation would finally determine the upper limit of output ratios.


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Enzyme immobilization is important for industrial applications,34 and previous studies have shown that the immobilization of enzymes on filamentous viruses may reduce their catalytic activities significantly.10 To investigate the activities of enzymes immobilized on HaCP nanotubes, nanoKAZ, a 19 kDa luciferase,35-36 was introduced into the HaCP nanotubes using a similar procedure to above (Figure 1B). nanoKAZ showed a higher incorporation ratio (nanoKAZ-HaCP:HaCP=1:2.8), compared with EGFP (EGFP-HaCP:HaCP=1:4.1), based on Western blotting analysis (Figure 2F, and Figure S5B). Immobilized nanoKAZ catalyzed the oxidation of coelenterazine, showing ~4-fold higher activities than free nanoKAZ (Figure 2G), possibly due to favorable alternations in the enzymatic system discussed before.34 In addition, immobilized nanoKAZ on the hybrid nanotubes retained ~90% activity (Figure 2G) after storage at 4 °C for 5 days, demonstrating that HaCP nanotubes are a stable scaffold for enzyme immobilization. Together these results show that HaCP nanotubes can provide efficient and stable bio-templates for the fabrication of complex hybrid nanostructures with precise control.

Cryo-EM studies on flexible HaCP nanotubes Compared with relatively rigid rod-shaped TMV, flexible filamentous viruses have shown improved capacity for loading larger functional moieties as partly decorated scaffolds.10 This is also the case for our assembled HaCP nanotubes. A reasonable explanation for this is that the detrimental modifications on viral CP could be buffered by soft bio-materials, and thus the accumulation of local assembly errors would be insufficient to disrupt nanotube formation. To design better long-range soft nanotubes, it is important to first know the general rules governing the assembly of these naturally occurring materials. Structural studies on flexible filamentous


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viruses have proven challenging,37 while HaCP nanotubes provide us with a new opportunity to study protein-only soft nanotubes. We investigated the assembly of HaCP nanotubes in their native state using cryo-electron microscopy (cryo-EM). Cryo-EM images showed that the tubes had a relatively smooth and uniform sidewall, which was characterized as two parallel dotted lines (Figure 3A–D), suggesting significant improvement in the structural signals compared with the distorted outlines of the negatively stained HaCP tubes. Some tubes displayed short-range straightness in the images (Figure 3A), but the majority exhibited some degree of curvature (Figure 3B). Strikingly, two slightly different morphological forms were observed in the samples (Figure 3C): narrow and wide tubes (hereafter referred to as N-tubes and W-tubes, respectively). There were many more N-tubes (~90%) in the preparations. Some N-tubes showed end-to-end fusion with W-tubes (Figure 3D), suggesting morphological transitions could happen during the assembly. For W-tubes, clear striation densities were found across the sidewall. In contrast, in the cryo-EM images N-tubes had no clear banding pattern, despite some similar but shorter striation stretches between the sidewall densities.


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Figure 3. Analysis of 2D class averaging from cryo-EM images of HaCP nanotubes. A–D) Cryo-EM images of selective HaCP nanotubes embedded in a thin layer of vitreous ice, recorded at a defocus range of 1–2.5 µm. A. a straight tube; B. a bent tube; C. an N-tube (unfilled arrow) and a W-tube (solid arrow); D. a tube consisting of a narrow domain (unfilled arrow) and a wide domain (solid arrow), with a transition region indicated by a white triangle. Scale bar: 50 nm. E) Gallery view of 2D class averages of extracted segment of N-tubes, created with a box size of 400 pixels and a round mask of 360 pixels diameter. The averages are horizontally aligned in the field and the numbers of segments for averaging are labeled in the lower left corner. F) 2D profiles for sidewall densities along a horizontally-oriented N-tube (left) or W-tube (right) class average. Red profile lines are for the upper sidewall and black profile lines are for the lower


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counterpart. Only the central 300 pixels are shown for calculations. Note the repeating pattern of N-tubes for every third helical pitch (green dashed lines).

The flexibility of HaCP nanotubes complicated direct applications of helical symmetry, and thus short but rather straight segments of nanotubes were extracted for single-particle-like analysis.38 The segments from N-tubes were grouped into distinct 2D classes (Figure 3E, and Figure S6A), while the segments from W-tubes could be grouped into only one class (Figure 3F, right). Notably, the 2D profiles of sidewall densities in N-tubes showed a reversed distribution for the opposite sidewall densities (Figure 3F, left), however a similar distribution pattern was observed for W-tubes (Figure 3F, right). The 2D class average of W-tubes bears significant resemblance to the characteristic projection of a cylinder consisting of stacked rings or disks, while the averages of N-tubes are reminiscent of the projections of a helix (Figure S7). In addition, N-tubes differ from W-tubes by displaying a repeating pattern at every third helical pitch for all class averages (Figure 3E–F), suggesting there is possible short-range symmetry along the axis of N-tubes. Two independent 3D maps were reconstructed for N-tubes and W-tubes at a resolution of 14 Å (based on a total of 17,923 segments, Table S1) and 21 Å (only 551 straight segments were used), respectively (Figure 4, and Figure S8). The solved 3D models showed characteristic surface grooves, which are consistent with the observed helical patterns in negatively stained nanotubes (Figure S8B, and Figure S9A). N-tubes are organized as a 1-start helix, in which a continuous helical line wraps around with 111⁄3 subunits per turn. In contrast, for W-tubes, stable lateral interactions between 12 subunits lead to them being arranged as ring-shaped building blocks,


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from which the elongated cylindrical structure is built. For helical reconstructions, W-tubes can also be viewed as 12 helices arranged in C12 rotational symmetry. Both N-tubes and W-tubes show an axial periodicity of ~10 nm (Figure 4A, and Figure 4C). In W-tubes, two rings are stacked 34.7 Å apart, which is equal to the pitch height of the 1-start helix in N-tubes. For Ntubes, every 34 subunits in 3 turns share a repeating pattern along the cylinder axis. For W-tubes, each ring of 12 subunits rotates 11.25° relative to the neighboring ring, and a total of 36 subunits in three adjacent rings could be viewed as an approximate repeating unit (although 10° rotation would be the ideal case).

Figure 4. Cryo-EM reconstructions of N-tubes and W-tubes. A) 3D density map of N-tubes at a resolution of 14 Å shows the symmetry of the tube. 12 adjacent subunits along the one-start helical path are color-coded (green for the first subunit, blue for the 2nd to 11th ones, and red for the 12th one). Along the tube axis, subunits on N-tubes repeat exactly in 3 turns (denoted by R). The pitch of 1-start helix is denoted by P. Note: since no atomic structures for HaCP are


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available, precise molecular boundaries cannot be defined at this point. The subunit here represents a minimal symmetry unit for helical reconstructions only. B) Two different views of the 12-subunit turn are shown separately, from left (0°) to right (45°). Two arrows mark the inter-layer contact surface for one subunit in an enlarged view. C, D) 3D density map of W-tubes at a resolution of 21 Å showing different features compared with N-tubes. Subunits on W-tubes repeat roughly for every three consecutive rings (denoted by R*).

If the helical system could relax (N-tubes) or pressure out (W-tubes), the diameter of the helix would change from 25 nm to 28 nm (Figure 5A, and Figure S6). The ratio of inner/outer diameter remains the same (64%) for the two tube types. And thus, theoretically, HaCP N-tubes could provide 439 m2/g of exterior surface area (281 m2/g of interior surface) for loading of functional molecules, and W-tubes could provide ~6% more exposed surface (464 m2/g of outer surface and 298 m2/g of inner surface). In contrast, RNA-directed assembly of TMV CP gives rise to a much thicker tube with a lower inner/outer diameter ratio (22%) 39 (for comparison, see Figure 5A, and Table S2). In structural engineering, the area moment of inertia is a geometrical property of plane sections of columns, which provides insight into the resistance of a column to bending. However, the calculated area moments of inertia for TMV CP nanotubes, HaCP Ntubes, and W-tubes, are 5,140 nm4, 15,958 nm4, and 25,019 nm4, respectively, suggesting that HaCP nanotubes are physically more resistant to external forces. Therefore, the flexibility of HaCP nanotubes is likely to be related to their intrinsic softness from elastic inter-subunit interactions.


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To better visualize lateral protein-protein interactions, 12 subunits along the 1-start helix of the N-tube, and 12 subunits in a ring of the W-tube were extracted for comparison (Figure 4B, and Figure 4D). Density continuities of lateral subunits can be observed even at the 3σ contour level for both tube types (for comparison of different σ, see Figure S9). In contrast, substantial lowdensity regions (corresponding to the holes in the tube wall in the 3σ maps) reduce the interface between two neighboring layers (i.e. turns of N-tube or rings of W-tube), resulting in only two restricted contacting regions within one subunit (Figure 4B, and Figure 4D). These observations indicate that lateral interactions might serve as dominant stabilizing forces for tube assembly, while weaker inter-layer interactions might contribute to the structural flexibility. Clear striation densities on the W-tubes provide a unique opportunity for analysis of inter-layer distance for single nanotubes (Figure 5B). Although only two stable conformational states (N-tubes and Wtubes) had been structurally calculated from the data, we expect that there are other metastable conformations (Figure 5C). The lattice-like surface can be compressed or expanded by adjusting inter-layer distance assuming relative distances between lateral subunits are unchanged. These slight differences could cumulatively give rise to varied axial spacings for different regions on the tube (Figure 5B), thereby altering local curvature without changing the number of subunits per layer. This flexibility provides HaCP nanotubes substantial spatial freedom, and could therefore be used for loading different functional modules without sacrificing the capacity for self-assembly.


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Figure 5. Structural comparison of TMV and HaCP nanotubes. A) Diagrams showing the arrangement of TMV CP and HaCP. Roughly elongated TMV CP molecules extend radially outward from the central channel (left), while the HaCP subunits in the thinner sidewall are largely lined tangentially along the one-start helix of N-tubes (middle) or along the ring of Wtubes (right). B) A curved W-tube showing characteristic striation densities along the tube axis. C) Enlarged view of 2 neighboring striation densities. Bending of the W-tube results in varied inter-layer distances from 34.0 Å to 35.4 Å (see Figure S11), which could be achieved by stretching or compressing the lattice-like surface consisting of HaCP subunits. Note that the


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average inter-ring distance is 34.7 Å for W-tubes, consistent with the cryo-EM reconstruction of W-tubes.

Cryo-EM studies were also performed for mKate-HaCP/HaCP hybrid nanotubes. As expected, there was no density layer corresponding to mKate because mKate is distributed on 20% of the exterior surface of the particle rather than forming a continuous layer (see above). Compared to W-tubes, N-tubes appeared still the dominant species in the hybrid nanotubes. The uneven introduction of heterogeneous protein components onto the protein nanotube also lowers the resolution of the map. However, based on 2D class averaging and a 3D reconstruction at 37-Å resolution (Figure S10), the overall structure of the hybrid N-tube remains the same as the HaCPonly nanotube. The relatively stable configuration of hybrid HaCP nanotubes in an aqueous system provides a uniform platform for loading functional groups, and their stability in other organic solvents would be also investigated next for more potential applications. Baculovirus CP in the wild-type particles, arranged into tube-like structures as stacked rings running parallel to the cylinder axis, as suggested by a low-resolution structural model based on early studies of empty nucleocapsids of Spodoptera litura granulosis virus (SpliGV, a Betabaculovirus).40 Our W-tubes share the same symmetrical and periodic arrangements as observed in empty nucleocapsids of baculovirus SpliGV, suggesting the recombinant HaCP expressed and purified from E. coli behaves in a fashion similar to naturally occurring HaCP. The building subunits in both N-tubes and W-tubes could be HaCP monomers as suggested by comparison of their density volume with 26 kDa EGFP (Figure S9), which is also consistent with structural analysis of SpliGV nucleocapsids. It has been proposed that the empty capsid sheath is


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pre-formed before the baculovirus genome is packaged into it.41 However, our results are less reconciled with this model. In the absence of other viral components, N-tubes (~90% in our preparations, see above), not W-tubes, are likely to be more readily formed. Therefore, it would be reasonable to expect the formation/selection of the capsid shell in vivo is coupled to other molecular events, such as the encapsulation of the genome or asymmetrical incorporation of other minor capsid components. Flexible capsid at this stage could provide a solution to accommodate a number of minor capsid proteins. The flexibility of baculovirus capsid needs further investigation.

CONCLUSIONS There are multiple properties that define ideal CP-based materials in terms of incorporation of functional modules: high incorporation efficiency, capacity to load large active molecules, and reasonable economic availability. Our results provide proof-of-principle demonstration that HaCP nanotubes could be used as an efficient hybrid platform for presenting multiple functional groups simultaneously, in near physiological conditions. Presentation of full-length proteins on the HaCP nanotubes also demonstrated that these materials could provide a large capacity for loading heterogeneous functionalities. The E. coli expression system reduces the cost and the complexity of synthesis and post-synthesis modifications, making heterogeneous formulations a possibility. Synthesizing new bio-nanotubes for materials studies based on structurally well-known molecules, such as cyclic assemblies of SP1,42 GroEL,43 Hcp1,44 and TRAP,45 has been a hot topic for research. As natural intermediates, HaCP nanotubes are resistant to the complex cellular


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environment and could provide a more bio-friendly alternative to these designed nanotubes. In addition, cryo-EM studies suggested that local tunable inter-subunit interactions give rise to the flexibility of HaCP nanotubes, resulting in promising soft materials as bio-templates for functionalization. Nature has provided an enormous amount of functional materials and we still have a long way to go in reproducing them to our advantage in a controlled way.

MATERIALS AND METHODS Gene Cloning. The full-length HaCP gene was amplified from the baculovirus Helicoverpa armigera nucleopolyhedrovirus (HearNPV) genome (GenBank Accession No. AF271059). The PCR product was cloned into the BamHI and XhoI sites of expression plasmid pGEX-6P-1 (Pharmacia Biotech). The resulting fusion protein contained an N-terminal GST tag, a 3C cleavage site and C-terminal HaCP (GST-HaCP). Genes of fluorescent proteins (EGFP, DsRed, mTagBFP and mKate) and luciferase (nanoKAZ) were codon-optimized for the E. coli expression system and then synthesized by Sangon (Shanghai). These genes were inserted into GST-HaCP fusion protein between the 3C cleavage site and C-terminal HaCP sequence (Figure S1B). To increase the cleavage activity of 3C protease, a short stretch of two or five glycine residues was introduced between the cleavage site and inserted fragments. The resulting protein products for nanotube assembly were HaCP, EGFP-HaCP, DsRed-HaCP, mTagBFP-HaCP, mKate-HaCP, and nanoKAZ-HaCP. Assembly of HaCP Nanotubes. Recombinant plasmids were transformed into BL21 (DE3) competent E. coli cells. For expression, a single colony was selected and grown at 37 °C


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overnight in 100 ml of LB medium containing 50 µg/ml of ampicillin. The pre-cultures were then inoculated (1%) in LB medium. The culture was incubated at 37 °C until an OD600 of 0.6– 0.8 was reached. Expression of the protein was induced by the addition of isopropyl β-Dthiogalactopyranoside (IPTG) to a final concentration of 0.5 mM and the cells were grown for another 20 h at 16 °C. The cells were harvested by centrifugation at 4,000 g for 30 minutes at 4 °C, suspended in 1×phosphate-buffered saline (PBS, pH 7.4) and pelleted again by centrifugation. The pellet was re-suspended in ice-cold assembly buffer containing 1×PBS (pH 7.4), 5% (v/v) glycerol, 0.01% Triton X-100, and 5 mM DTT. The cells were lysed by sonication, and the lysate was then centrifuged for 80 min at 10,000 g at 4 °C. The supernatant was filtered using a 0.45 µm filtering unit (Millipore) and loaded onto a gravity column containing glutathione agarose resin (GE Healthcare). The resin was washed with ~50 bed-volumes of the assembly buffer. HaCP was freed from GST-HaCP by incubation with GST-3C protease (a fusion protein of N-terminal GST and C-terminal human rhinovirus B14 strain 3C protease, prepared in our lab) and assembled into nanotubes at 4 °C overnight in the assembly buffer. As a control, 500 U deoxyribonuclease I (DNase I, C&M Biolabs) with 40 µM MgCl2 (Sigma) was applied to the GST column before adding GST-3C to remove any remaining DNA. Hybrid HaCP nanotubes were prepared in a similar way. First, 20% of the cell cultures were used for affinity purification of each GST-tagged protein (i.e. GST-HaCP, GST-EGFP-HaCP, etc.) according to the manufacturer's protocol. The yield of each protein was then determined by the Bradford method using bovine serum albumin (BSA) as a standard (Thermo Fisher Scientific). For the EGFP-HaCP/HaCP hybrid assembly, cell cultures expressing EGFP-HaCP and HaCP were mixed with a corresponding molar ratio of 1:2 before being lysed by sonication, followed by GST affinity purification and 3C digestion as described above. Other assembly formulations


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were mKate-HaCP/HaCP (1:2, 1:4, 1:8, and 1:16), EGFP-HaCP/DsRed-HaCP/mTagBFPHaCP/HaCP (1:1:1:6), nanoKAZ-HaCP/HaCP (1:2), EGFP-HaCP/nanoKAZ-HaCP/HaCP (1:1:4). Purification of Nanotubes by SEC. After 3C digestion the nanotube samples were concentrated using Amicon filter units with a 30 kDa molecular weight cutoff. Further purification was carried out using SEC on a pre-packed Sephacryl S-500 High Resolution column (GE Healthcare) preequilibrated in 1×PBS (pH 7.4). The first peak eluted by PBS was identified as HaCP nanotubes and concentrated to 0.5 mg/ml and stored at 4 °C. The identity of the protein from the second peak was determined using peptide mass fingerprinting (PMF) techniques as described previously.20 In brief, mass spectrum data of the peptide mixture from trypsin digestion were analyzed by a Mascot search against the NCBI nr database. The matched protein was 60 kDa molecular chaperone GroEL of E. coli (GenBank Accession No. WP_000729117). Immunoblotting and Molar Ratio Estimation. SDS–PAGE analysis was carried out under reducing conditions using a 12% separation gel. SDS-PAGE gels were transferred by semi-dry transfer apparatus (Bio-Rad) to nitrocellulose membranes. Membranes were blocked using 5% skim milk in PBS. Rabbit anti-HaCP antibodies (prepared by ourselves) were used for probing HaCP epitopes and were detected with peroxidase-coupled secondary antibodies. To estimate the molar ratio of X-HaCP to HaCP (X stands for functional moieties) in the nanotubes, bands on nitrocellulose membranes were scanned and integrated using the Quantity One package (Bio-Rad) according to the user manual. The ratios were estimated by assuming that coupled peroxidase is linearly proportional to the amount of proteins on the membrane.


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Transmission Electron Microscopy (TEM). TEM was used to visualize negatively stained HaCP nanotubes and other hybrid nanotubes. ~5 µl of sample (sufficient to cover the grid surface) was placed on a glow-discharged EM grid with continuous carbon film for 60 sec. Excess solution was removed by touching the grid edge with filter paper. The grid was then rinsed with ten drops of ddH2O. 10 µl of 2% phosphotungstic acid (PTA) was then applied to the grid for 30 or 300 s. Excess stain was removed by touching the grid edge with filter paper. The grid was allowed to dry in air for 5 min. The grids were imaged by TEM using an FEI Tecnai G2 20 TWIN operating at 200 kV and an Olympus SIS Cantega G2 2K×2K CCD. HaCP Nanotubes at Acidic and Alkaline pH. Different pH buffers were prepared to test the stability of HaCP nanotubes: 100 mM citric acid-sodium citrate buffer (pH 3.0), 100 mM acetic acid-sodium acetate buffer (pH 4.0, 5.0, and 6.0), 100 mM Tris buffer (pH 8.0 and 9.0), 100 mM CAPS buffer (pH 10.0 and 11.0). HaCP nanotubes (0.5 mg/ml, in 1×PBS) were diluted with different pH buffers at a ratio of 1:9, and then incubated at 4 °C for 12 h. The samples were then subjected to negative staining and examined using TEM as described above. Immuno-electron Microscopy (IEM). IEM was performed to confirm that EGFP-HaCP had coassembled with HaCP into nanotubes. 5 µl of 0.1 mg/ml sample was loaded onto a glowdischarged grid for 30 min at RT. The grid was washed three times with PBS, and then incubated in PBS containing 5% skim milk for 15 min to block nonspecific binding. Excess buffer was removed using filter paper. The grid was incubated with rabbit anti-EGFP polyclonal antibody (Proteintech) diluted 1:50 in PBS for 60 min and washed five times with PBS at RT. Excess PBS was removed using filter paper. The grid was then labeled with 10 nm gold particles, which were conjugated to goat anti-rabbit IgG (Bioss) at a dilution of 1:50 for 60 min at RT. The grid was washed six times with PBS and three times with ddH2O. Excess ddH2O was removed with filter


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paper. The same negative staining procedure as described above was then performed to increase signal. The distribution of nanogold particles on EGFP-HaCP/HaCP nanotubes was examined by TEM. Fluorescence Microscopy. To minimize the movement of HaCP nanotubes, glass bottom dishes (NEST) were coated with 50 µl of 0.01% poly-L-lysine (Sigma) and allowed to dry in air at 37 °C overnight. 0.05 mg/ml HaCP nanotube samples were then added to the pre-treated dishes. The samples were examined immediately with fluorescence microscopy. Images of EGFPHaCP/HaCP nanotubes and EGFP-HaCP/DsRed-HaCP/mTagBFP-HaCP/HaCP nanotubes were collected using NIS-Elements software (Nikon) on a Nikon A1 MP confocal microscope with an oil immersion 60× Plan Apo 1.40 NA Nikon objective. DAPI filters were used to visualize mTagBFP-HaCP, FITC filters were used to visualize EGFP-HaCP, and TRITC filters were used to visualize DsRed-HaCP. Grayscale nanotube images were colored red, green, or blue according to the filter cube set used, and then merged using NIS-Elements. Enzymatic Assay for nanoKAZ. Before enzymatic assay for nanoKAZ, nanoKAZ-HaCP in nanoKAZ-HaCP/HaCP nanotubes, and free nanoKAZ were quantified by SDS-PAGE with BSA as an internal standard. The reaction mixture (100 µL) contained 1 µg coelenterazine, 30 mM Tris-HCl (pH 7.6), and 10 mM EDTA. The luminescence activity was determined using the luminescence mode of an EnSpire Multimode Plate Reader (PerkinElmer). The luminescence reaction was triggered by the addition of 10 µL of diluted nanoKAZ-HaCP/HaCP nanotubes (corresponding to 0.005 nmol of nanoKAZ) into the reaction mixture, and the luminescence intensity was recorded in 0.1 s intervals for 600 s. Control assays were performed using 0.005 nmol of free nanoKAZ enzyme.


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Cryo-electron Microscopy. A 3 µl aliquot of HaCP nanotubes (0.5 mg/ml, in PBS) was applied to a glow-discharged GiG R2/1 holey carbon grid. The grid was then blotted using filter paper and plunge-frozen in liquid ethane using a Vitrobot plunging system (FEI). Imaging was performed using an FEI Titan Krios (Zhejiang University) operating at 300 kV and a Gatan K2 Summit direct electron detector via the SerialEM software package.46 A total of 379 movies (each containing 24 frames of 3710×3838 pixels) of HaCP-only nanotubes and 205 movies of mKate-HaCP/HaCP hybrid nanotubes, were recorded at 1.014 Å per pixel with a defocus range of 1–2.5 µm. The total dose for each movie was less than 40 electrons per Å2. For each movie, the frames were aligned and a single averaged micrograph was generated by MotionCor.47 Particle picking, 2D reference-free classification, and 3D classification and refinements were performed using Relion 2.0.38 For N-tubes, manual and template-based particle picking strategies were carried out and particles were then extracted along curved nanotubes with a box size of 400 pixels with a 35-pixel shift between adjacent boxes. CTF estimation was carried out with the CTFFIND4 package

48

in the Relion workflow. 25 rounds of reference-free 2D classification

were performed and particles in the best 34 classes (total 100 classes) were used for subsequent calculations. To determine helical symmetry for the N-tubes, a single-particle-like analysis was used, instead of traditional Fourier-Bessel indexing. A featureless tube was used as a starting model for 3D refinement, and the resulting asymmetrical model was used for estimation of twist and rise, by which a new model was generated for the final 3D refinements (Figure S8A). For 3D analysis of W-tubes, 10 micrographs with clear features of W-tubes were manually selected and particles were picked with the same procedure as used for N-tubes. We used a rise range of 33.9– 34.9 Å (inter-layer distance was 34.4 Å based on 2D averaging) and a twist range of 9°–15° as searching parameters for 3D refinement. Since there could only be an integral number of


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subunits in a ring for W-tubes, different rotation symmetries (C10, C11, C12, C13, C14, etc.) were tested. A reconstruction for C12 symmetry converged after 20 iterations with clear structural features. Handedness of N-tubes and W-tubes was determined by visual comparison of the left- or right-hand reconstructions with the helical pattern on the negatively stained tubes (Figure S8B). Using the gold-standard Fourier shell correlation (FSC) = 0.143 cutoff, the map resolution was estimated to be 14 Å and 21 Å for N- and W-tubes, respectively (Figure S8C and Figure S8D). Segmentation of density maps was performed with Chimera.49 The area moment of inertia for nanotubes was calculated using the Equation (1). The area moment of inertia for

I = nanotubes was calculated using the following equation:

π

(D 64

4

−d4

)

,

where D = outer diameter and d = inner diameter. All of the images of 3D reconstructions were prepared with Chimera. Maps were restored in EMDataBank as accession numbers of EMD6933 (N-tubes) and EMD-6936 (W-tubes). Statistics. P values were calculated using two-tailed, unpaired Student’s t tests in Graphpad Prism. The statistical tests used are indicated in the figure legends, and P values < 0.05 were considered significant.

ASSOCIATED CONTENT Supporting Information. The following files are available free of charge. Additional figures and tables give details on the assembly, purification and cryo-EM analysis of HaCP nanotubes (PDF).


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AUTHOR INFORMATION Corresponding Authors *E-mail: [email protected]. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by funds from the National Natural Science Foundation of China (31770169) and the Key Research Projects of Frontier Science, Chinese Academy of Sciences (QYZDJ-SSW-SMC021). The cryo-EM data collection was conducted at the Center of CryoElectron Microscopy at Zhejiang University. We thank Prof. Xing Zhang and Dr. Shenghai Chang from Zhejiang University for assistance with the cryo-EM. We thank Drs. Fang Li and Wei Zhang from the University of Minnesota for initial discussion. We thank the Center for Instrumental Analysis and Metrology of Wuhan Institute of Virology, CAS, for the assistance in acquiring confocal and electron microscope images (Dr. Ding Gao). We thank Dr. Sarah Dodds for editing the English text of a draft of this manuscript.

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Graphic Table of Contents


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Controllable Assembly of Flexible Protein Nanotubes for Loading

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