Design of Modular Polyhydroxyalkanoate Scaffolds for Protein

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Design of modular polyhydroxyalkanoate scaffolds for protein immobilization by directed ligation Jin Xiang Wong, and Bernd H. A. Rehm Biomacromolecules, Just Accepted Manuscript • Publication Date (Web): 10 Sep 2018 Downloaded from http://pubs.acs.org on September 10, 2018

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Biomacromolecules

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Design of modular polyhydroxyalkanoate scaffolds

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for protein immobilization by directed ligation

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Jin Xiang Wonga,c, Bernd H. A. Rehmb*

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a

Institute of Fundamental Sciences, Massey University, Private Bag 11222, Palmerston North,

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New Zealand b

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Centre for Cell Factories and Biopolymers, Griffith Institute for Drug Discovery, Griffith University, Don Young Road, Nathan, QLD, Australia

c

MacDiarmid Institute of Advanced Materials and Nanotechnology, Victoria University of Wellington, Wellington 6140, New Zealand

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ACS Paragon Plus Environment

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Biomacromolecules 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Abstract

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In vivo assembled polyhydroxyalkanoate (PHA) particles have been successfully bioengineered to

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display foreign protein functions towards high-value applications in medicine and industry. To

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further expand the design space of PHA particles towards immobilization of various functional

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proteins, we developed a tunable modular protein immobilization method implementing the

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SpyCatcher/SpyTag chemistry. We successfully displayed the SpyCatcher protein using

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translational fusion with the Ralstonia eutropha PHA synthase (PhaC). The SpyCatcher domain

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displayed on the surface of PHA particles was accessible for cross-linker-free ligation with

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SpyTag-bearing proteins. We demonstrated tunable protein immobilization of various SpyTagged

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proteins on SpyCatcher-PHA particles, which ultimately enabled assembly of multiple proteins

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coating the surface of PHA particles. Overall, the functionality, stability, and reusability of proteins

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immobilized to SpyCatcher-PHA particles were either retained or enhanced in comparison to the

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soluble forms. This modular platform can be implemented as a generic tool for protein

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immobilization in an array of applications.

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Keywords: Polyhydroxyalkanoates, protein immobilization, enzyme immobilization, modular,

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SpyCatcher-SpyTag chemistry


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Biomacromolecules

INTRODUCTION

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Protein immobilization techniques have long been recognized as useful tools for real-world uses

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in biomedical and industrial sectors. Immobilization of proteins to the surface of support materials

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allows the design of favorable microenvironments to achieve optimum performance. Improvement

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in both the stability and functionality of immobilized proteins has been reported by taking

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advantage of nonspecific interactions between different proteins and supporting materials

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enhancing functional conformation and orientation.1-5 However, adverse effects were observed

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when interactions with surfaces unfavorably impacted on orientation and conformation of

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immobilized proteins.6-8 The close-proximity between proteins upon immobilization improves the

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functional performance and stability, as a result of the macromolecular crowding.9-11 Several

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studies reported an improvement of Vmax of enzymes in crowded microenvironments due to an

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increase in effective concentration of enzymes.10,

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achieved by macromolecular crowding, where the protein unfolding equilibrium can be shifted in

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the direction leading to the formation of thermodynamically rigid proteins, had been

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demonstrated.9, 13 Minton developed a statistical thermodynamic model to estimate the stability of

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globular proteins under temperature stress and in the presence of chaotropic agents with respect to

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the excluded volume effect and he predicted that crowding of stable globular proteins sufficiently

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improved both the thermal and chaotropic stability.14 Nevertheless, some authors also noted

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reduced protein functionality due excessive crowding of respective proteins on solid supports.15-16

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Macromolecular crowding has also been linked to the formation of protein aggregation, which

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leads to strong inhibition of protein functionality.17-18 Moreover, clustering different proteins in

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the immobilized form on scaffolds also allows biomimicry of co-localized proteins as found in

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natural multiprotein complexes, where many attempts have been devoted to reconstitute such

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Also, greater stability of proteins can be


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capabilities.19-21 Protein immobilization is also crucial for industrial applications, especially in the

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case of enzymes, where it stabilizes enzymes and allows their use in continuous processing

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providing economic advantages related to recovery and reuse of enzymes.22

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Polyhydroxyalkanoates (PHAs) have been recently considered as support materials for in vivo

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protein immobilization. PHAs are deposited inside the bacterial cell as spherical polyester

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inclusions and are naturally produced under unbalanced nutrient conditions. Bacterial productions

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strains can be developed that self-assemble PHAs to form shell-core structures, where the surface

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can be functionalized by protein engineering of PHA-binding proteins and chemical means.23

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Recently, we showed that formation of such PHA particles inside bacterial cells can be tailored for

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surface display of a range of protein functions by using recombinant DNA technology, leading to

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the development of the PHA particle technology as a versatile platform for protein immobilization

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and display.24-30 This new technology is based on the translational fusion of functional proteins of

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interest to the N- and/or C-terminus of a PHA synthase (PhaC), which results in in vivo self-

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assembly of PHA particles displaying such functional proteins.31-32 PhaC itself catalyzes PHA

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synthesis and remains covalently attached to the surface of the PHA particles.33-34 While the use

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of PhaC as anchoring domain represents an efficient way of immobilizing proteins to PHA

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particles, it is depending on the biological complexity of the bacterial production strain inherently

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limiting control over PHA particle formation.24, 30 Misfolding, low-density surface display and

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potential failure in achieving multifunctionality indicate some limitations of the use of the PHA

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particle technology. In addition, it will be challenging to control the ratio of certain functionalities

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using the PHA particle technology where assembly of functional PHA particles occurs inside the

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bacterial cell. Also, in some cases high-value functional proteins (e.g., eukaryotic therapeutic

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proteins) might require a production hosts catering for protein folding pathways as well as post-


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Biomacromolecules

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translational modifications for proper folding and functionality. However, these production hosts

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are suboptimal for cost-effective production of the PHA carrier material.28-29 Hence, in some cases,

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it might be advantageous to separately produce PHA particles and proteins of interest under their

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respective optimum conditions followed by in vitro chemical conjugation of the protein of interest

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to the PHA particle surface. However, these chemical modifications are often laborious, potentially

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disrupt the native functionality of proteins as well as lead to random protein orientation.35 To

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address these issues, the site-specific protein ligation system, SpyCatcher/SpyTag chemistry36,

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derived from CnaB2 domain from the fibronectin-binding protein (FbaB) found in Streptococcus

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pyogenes, might offer an efficient alternative for oriented functional immobilization of proteins to

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PHA particles.

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The SpyCatcher/SpyTag chemistry offers a very promising protein ligation tool as it can be

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carried out under a wide range temperatures (4°C to 37°C), pH values (5 to 8), selection of buffers

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(anion, cation or non-ionic) and it does not require the use of chemical cross-linkers or enzymes.

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The SpyCatcher is a small protein comprising 116 amino acid residues. It is able to spontaneously

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form an isopeptide bond with a 13 amino acid residue short peptide (SpyTag), by mixing these

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two components without the need of additional enzymes or chemicals.37 In recent years, there has

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been growing interest in utilizing the SpyCatcher/SpyTag chemistry in designing different modular

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scaffolding systems for protein immobilization, and surface functionalization, such as virus-like

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particles, various protein scaffolds/cages, gold nanoparticles, silica supports for potential

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applications in vaccination, bio-imaging, and synthetic biology.38-47

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The aim of this study was to design a generic modular immobilization system for proteins by

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merging the PHA surface display technology with the versatile SpyCatcher/SpyTag chemistry. We

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aim to display the SpyCatcher protein at high density on the surface of in vivo self-assembled PHA


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particles via translational fusion of SpyCatcher to the N- or C-terminus of PhaC. The SpyCatcher

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will serve as a covalent ligation site for SpyTagged functional proteins. To demonstrate the broad

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applicability of this new approach we will design and produce several SpyTagged proteins,

4

representing diverse functional categories, for site-specific ligation to SpyCatcher coated PHA

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particles. This study will also investigate the tunability of the modular system in order to achieve

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multiple protein functions. Functionality, stability, and reusability of resulting functional PHA

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particles will be analyzed.

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EXPERIMENTAL SECTION

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Bacterial strains, genetic manipulation, growth conditions. All the bacterial strains,

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plasmids, and primers used in the current study are listed in Table S1, Table S2, and Table S3

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respectively, which can be found in the supporting material. The SpyCatcher encoding DNA was

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synthesized by Genscript (Piscataway, USA) and primers were ordered from Integrated DNA

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Technologies (San Diego, USA). General DNA isolation, manipulation, and cloning procedures

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were performed as described elsewhere.48 For plasmid propagation and cloning, E. coli XL1-Blue

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(Stratagene, La Jolla, USA) was grown overnight (16 h) in Luria Bertani (Lennox) medium (pH

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7.5) at 37°C and 200 rpm. When needed, ampicillin (100 μg/mL) and chloramphenicol (50 μg/mL)

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were added. DNA sequences of the newly constructed plasmids were sequenced by Massey

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Genome Service (Palmerston North, New Zealand).

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Newly constructed plasmids used for this study were transformed into competent E. coli

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BL21(DE3) cells (Invitrogen, Carlsbad, USA), and competent E. coli BL21(DE3) cells harboring

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plasmid pMCS69 for production of soluble free proteins and PHA particles, respectively. Plasmid


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Biomacromolecules

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pMCS69 present in the latter strain enables the production of the precursor R-3-hydroxybutryl-

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coenzyme A (CoA), which is required for PHA synthesis. Detailed plasmid construction strategies

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can be found in the supporting material.

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Polyhydroxyalkanoate (PHA) particle production and isolation. Overnight culture of the

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respective E. coli BL21 (DE3) strains were diluted 1:100 into fresh Luria Bertani, Lennox (LB-

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Lennox) medium containing ampicillin and chloramphenicol supplemented with 1% (wt/vol)

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glucose. The culture medium was cultivated at 37°C and 200 rpm until an OD600 value of 0.6 - 0.8

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was achieved. PHA particle production was induced by addition of 1 mM of isopropyl β-D-1-

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thiogalactopyranoside (IPTG) into the cultures. Cultures were grown 48 h at 25°C. The cell pellets

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harvested were (8,000 g at 4°C for 20 min) washed with 10 mM Tris-HCl (pH 7.5) once using a

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homogenizer (MICCRA D-9 45132, Müllheim, Germany) prior cell disruption. Cells were lysed

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as previously described and PHA particles were recovered by centrifugation (9,000 g at 4°C for

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20 min).49 Recovered PHA particles were then washed three times and resuspended in PHA

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particle storage buffer (50 mM Tris-HCl, 20% vol/vol ethanol, pH 7.5) and stored at 4°C for further

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use and analysis.

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Production and purification of soluble protein. Overnight culture of the respective E. coli

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BL21 (DE3) strains were diluted 1:100 into fresh LB-Lennox medium containing ampicillin and

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cultivated at 37°C and 200 rpm until an OD600 value of 0.6 - 0.8 was achieved. Protein production

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was induced by addition of 1 mM of IPTG to the cultures. Cultures were harvested after 24 h

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incubation at 30°C. The cell pellets harvested were (8,000 g at 4°C for 20 min) washed with 10

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mM Tris-HCl (pH 7.5) once using a homogenizer (MICCRA D-9 45132, Müllheim, Germany)

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prior cell disruption. Washed cell pellets were resuspended to 10% cell slurry in 1X protein lysis

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buffer (50 mM Tris-HCl, 300 mM NaCl, 10 mM imidazole, pH 7.5) and lysed by passing through


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a microfluidizer (M-110P, Microfluidics, Westwood, USA) at 1500 bar. After cell lysis, the whole

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cell lysate was centrifuged (9,500 g at 4°C for 1h) to discard the cellular debris. The supernatant

3

was filtered through a 0.22 μm cellulose acetate membrane filter (ReliaPrepTM, Ahlstrom-

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Munksjö, Helsinki, Finland) and the cleared supernatant was loaded through 5 mL Protino® Ni-

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NTA column (Macherey-Nagel, Düren, Germany) at 5 mL/min. The Ni-NTA column was washed

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with at least 5 column volumes of protein wash buffer (50 mM Tris-HCl, 300 mM NaCl, 50 mM

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imidazole, pH 7.5) to remove non-specifically bound proteins. The proteins were eluted in protein

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elution buffer (50 mM Tris-HCl, 300 mM NaCl, 500 mM imidazole, pH 7.5) with at least 5 column

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volumes. Eluted protein samples were concentrated and desalinated using centrifugal concentrator

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(Vivaspin® 20, GE Healthcare, Buckinghamshire, U.K.). Subsequently, concentrated samples

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were stored at 4°C for further use and analysis.

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Protein analysis. All fusion proteins were analyzed by sodium dodecyl sulfate–polyacrylamide

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gel electrophoresis (SDS-PAGE) as described elsewhere.50 Briefly, protein samples were

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separated in 10% (vol/vol) polyacrylamide separating gels with 4% (vol/vol) polyacrylamide

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stacking gels. The molecular mass of the samples was estimated using GangNam-STAINTM pre-

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stained protein ladder (iNtRON Biotechnology, Seongnam, South Korea). SDS-PAGE gels were

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stained with 0.05% (wt/vol) Coomassie brilliant blue R-250 dye, 50% (vol/vol) ethanol and 10%

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(vol/vol) acetic acid for 30 min, then destained in 50% (vol/vol) ethanol and 10% (vol/vol) acetic

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acid for 2 h. Images of polyacrylamide gel were taken using Gel Doc XR+ system (Bio-Rad

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Laboratories, Hercules, USA).

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Protein quantification. Protein concentration was determined by measuring the band intensity

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from SDS-PAGE gels for densitometric analysis using Image Lab 5.2.1 software (Bio-Rad


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Biomacromolecules

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Laboratories, Hercules, USA), and comparing the value to a standard curve prepared from known

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concentrations of bovine serum albumin (BSA) standard as described elsewhere.51

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Peptide mass fingerprinting. Purified protein bands from the SDS-PAGE gel were excised

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and subjected to tryptic in-gel digestion as described elsewhere.52 The resulting tryptic peptide

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samples were then analyzed by liquid chromatography-tandem mass spectrometry (LC-MS/MS).

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Immobilization of SpyTagged proteins onto SpyCatcher-PHA particles. Feasibility of

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immobilizing SpyTagged Bacillus licheniformis α-amylase (BLA), SpyTagged green fluorescent

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protein (GFP), and SpyTagged organophosphohydrolase (OpdA) onto SpyCatcher-PHA particles

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was tested by incubating different SpyTagged proteins with SpyCatcher-PHA particles in 50 mM

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Tris-HCl, pH 7.5 overnight at 4°C under constant shaking at SpyCatcher:SpyTag reactant ratio of

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3:1. The samples were washed three times with 50 mM Tris-HCl, pH 7.5 before analyzed by SDS-

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PAGE analysis.

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Optimization of SpyCatcher/SpyTag chemistry. To optimize the reactant ratio, SpyCatcher-

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PHA particles were mixed with different SpyTagged proteins at SpyCatcher:SpyTag reactant ratio

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of 3:1, 2:1, 1:1, 1:2 and 1:3 in 50 mM Tris-HCl, pH 7.5. The mixtures were incubated overnight

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at 4°C under constant shaking. Then samples were washed three times with 50 mM Tris-HCl, pH

17

7.5 before analyzed by SDS-PAGE analysis. Meanwhile, the reaction time course of the ligation

18

chemistry was determined by incubating different SpyTagged proteins with SpyCatcher-PHA

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particles at SpyCatcher:SpyTag reactant ratio of 2:1 with a total reaction time of 24 h. The samples

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collected at 1 h, 3 h, 6 h, 12 h and 24 h, were washed three times with 50 mM Tris-HCl, pH 7.5

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before evaluated by SDS-PAGE analysis.


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Assembly of the immobilized multiprotein complex using SpyCatcher-PHA particles. To

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construct a proof-of-concept immobilized multiprotein complex system using the SpyCatcher-

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PHA particle platform, SpyTagged BLA was first incubated with SpyCatcher-PHA particles at

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SpyCatcher:SpyTag reactant ratio of 3:1 in 50 mM Tris-HCl, pH 7.5 overnight at 4°C under

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constant shaking. Next, the samples were centrifuged at 15,000 g for 10 min, and unbound proteins

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in each sample were discarded. The samples were washed three times with 50 mM Tris-HCl, pH

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7.5 and 10 μL of samples were taken for verification of protein ligation, by SDS-PAGE analysis.

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Same procedures were repeated by replacing SpyTagged GFP and SpyTagged OpdA, at

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SpyCatcher:SpyTag reactant ratio of 3:1 and 4:1, respectively.

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Compositional analysis of PHA particles. Approximately 75 mg of lyophilized PHA particles

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was subjected to methanolysis as described elsewhere.53 The organic layer of all samples was

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recovered, filtered and further analyzed by gas chromatography-mass spectroscopy (GC/MS)

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using poly (R)-3-hydroxybutyric acid (PHB) as standard.

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Zeta potential measurement. Zeta potential of the PHA particles was determined by

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electrophoretic light scattering (ELS) coupled with phase analysis light scattering (PALS) using

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Zetasizer Nano ZS (Malvern Instruments, Malvern, U.K.). All the PHA particle samples were

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measured in 0.1% (wt/vol) of the wet particles in 50 mM Tris-HCl, pH 7.5 and the soluble protein

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samples were measured in 50 mM Tris-HCl, pH 7.5.

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PHA particle size distribution measurement. Particle size distribution of the PHA particles

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was determined by dynamic light scattering (DLS) analysis using the Mastersizer 3000 laser

21

diffraction particle size analyzer (Malvern Instruments, Malvern, U.K.). The PHA particle samples


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Biomacromolecules

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were prepared in 0.1% (wt/vol) of the wet particles in storage buffer (50 mM Tris-HCl, 20%

2

(vol/vol) ethanol, pH 7.5).

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Fluorescence screening, microscopy analysis, and fluorescence intensity measurement.

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Fluorescence intensity of soluble free and immobilized GFP in 50 mM Tris-HCl, pH 7.5 were

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evaluated. The samples were first screened visually by the Safe Imager™ 2.0 Blue-Light

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Transilluminator (Invitrogen, Carlsbad, USA) and image of fluorescing samples excited with blue

7

light at 470 nm. Fluorescence microscope images of the samples were taken using an Olympus

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BX51 fluorescent light microscope (Olympus Optical, Tokyo, Japan) at 100X magnification using

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MicroPublisher 5.0™ color CCD camera, QCapture Pro 6.0 application software. (QImaging,

10

Surrey, Canada). Fluorescence intensity of the samples was measured using FLUOstar Galaxy

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fluorimeter and Reader Control Software (BMG Labtech, Ortenberg, Germany) at excitation and

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emission wavelengths of 380 nm and 520 nm, respectively.

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Starch degradation screen and colorimetric assay for α-amylase. Enzymatic performance

14

of immobilized and soluble free BLA with appropriate controls was first qualitatively verified by

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using starch agar plates.54 Briefly, 1% starch agar was prepared by dissolving 1% (wt/vol) soluble

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starch and 1.5% (wt/vol) agar with 50 mM Tris-HCl, 300 mM NaCl buffer (pH 7.5) prior to

17

autoclaving. All the samples were incubated at 37°C up to 24 h on the surface of the starch agar

18

plates. After the rapid screening, immobilized and free BLA, with negative controls were

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quantitatively assayed in a modified reaction buffer (50 mM Tris-HCl, pH 7.5)54 for their

20

functionality by 3,5-dinitrosalicyclic acid colorimetric method, using an amylase assay kit

21

purchased (Abcam, Cambridge, U.K.). Nitrophenol was liberated as the hydrolysis of ethylidene-

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pNP-G7BLA by BLA proceeded, and the release of nitrophenol was monitored by

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ELx808™Absorbance Microplate Reader using Gen5 reader control 1.02.8 application software


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(BioTeK Instruments, Winooski, USA) at OD405 nm under room temperature (25°C) for up to 3

2

h at 2 min intervals.

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Organophosphohydrolase functionality assay. Enzymatic performance of the immobilized

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and soluble OpdA (50 mM Tris-HCl, pH 7.5) with negative controls, were assessed using assay

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mixture of 250 μM coumaphos, dissolved in a modified reaction buffer (50 mM Tris-HCl, 20%

6

(vol/vol) methanol, pH 7.5).55 Quantification of liberated chlorferon from coumaphos was

7

determined by FluoroMax®-4 and Jobin Yvon MicroMax 384 microwell-plate reader controlled

8

by FluoEssence™version 3.5 (HORIBA Scientific, Kyoto, Japan) at excitation and emission

9

wavelengths of 355 nm and 450 nm respectively. Samples were loaded in an assay mixture and

10

performed at room temperature (25°C) for up to 2 h at 10 min interval.

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Thermal stability. Functional proteins in both immobilized and soluble forms were pre-

12

incubated from 5°C to 95°C at a temperature interval of 10°C using AccuBlock™ Mini Compact

13

Dry Bath (Labnet International, Edison, USA) for 30 min. The resulting samples were then

14

subjected to their respective functional assays for 1 h.

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pH stability. Immobilized and soluble free proteins at varying pH values were pre-incubated

16

in the following solutions for 30 min at room temperature (25°C): pH 3 and 5 (50 mM sodium

17

acetate), pH 7 and pH 9 (50 mM Tris-HCl) and pH 11 (50 mM disodium hydrogen

18

orthophosphate). Vivaspin® 20 centrifugal concentrator (GE Healthcare, Buckinghamshire, U.K.)

19

was used to perform the buffer exchange for soluble free proteins. After that, the samples were

20

resuspended in their reaction buffers, respectively after pH treatment and assessed for their

21

functionality for 1 h.


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Biomacromolecules

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Reusability. Both immobilized and soluble forms of functional proteins of interest were

2

measured for reusability with their respective functional assays in five consecutive cycles at room

3

temperature (2 h each cycle for OpdA and GFP; 3 h each cycle for BLA). Immobilized protein

4

samples were centrifuged at 15,000 g for 10 min in a microcentrifuge at the end of the assessment

5

cycle. The supernatant was discarded, and the samples were resuspended in fresh reaction buffers

6

respectively. Vivaspin® 20 centrifugal concentrator (GE Healthcare, Buckinghamshire, U.K.) was

7

used to perform the buffer exchange for soluble free proteins at the end of the assay, and the protein

8

samples were diluted with fresh reaction buffers. This procedure was repeated for five cycles.

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RESULTS AND DISCUSSION

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Design and production of SpyCatcher-displaying PHA particles. To enable efficient

12

ligation of proteins without the need of chemical cross-linkers or enzymes, we designed and

13

produced PHA particles displaying the SpyCatcher domain for ligation with SpyTagged proteins

14

of interest, where a covalent isopeptide bond forms between a lysine residue (Lys) of SpyCatcher

15

domain and an aspartic acid residue (Asp) of SpyTag peptide (Figures 1A and 1B). We

16

successfully displayed SpyCatcher on the surface of PHA particles via surface-exposed PHA

17

synthase (PhaC)56, using translational fusion of SpyCatcher to the N- and C- terminus of PhaC as

18

confirmed by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) (Figure

19

1C) and by peptide fingerprinting analysis using liquid chromatography-tandem mass

20

spectrometry (LC-MS/MS) (Supporting Information Table S4). The molecular mass of

21

SpyCatcher-PhaC (SP) and PhaC-SpyCatcher (PS) fusion proteins are 68.4kDa and 69.1kDa,

22

respectively, and greater than that of wild-type PhaC (WT) of 55.5kDa. Placing the SpyCatcher


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protein at the N-terminus of PhaC significantly enhanced the production yields of SP fusion protein

2

per PHA particle mass. We successfully overproduced the SP fusion protein displayed on PHA

3

particles (SP-P) resulting in yields of 194 nmoles SpyCatcher per g wet PHA particles and this

4

was much higher than found for PS fusion protein displayed on PHA particles (PS-P) (Figure 1D).

5

Also, it has been demonstrated that the N-terminus fusion point of PhaC is located at a highly

6

variable surface-exposed region of the protein itself and proven to be not essential to the PhaC

7

activity.33, 57 In contrast, the C-terminus of PhaC is conserved and essential for PhaC activity and

8

was proposed to be attached to the inner hydrophobic core of PHA particles, thus potentially

9

affecting the surface exposure of any C-terminally fused domains.32 Therefore and due to the high-

10

density display of exposed SpyCatcher domains, the SP-P was selected to demonstrate the proof-

11

of-concept for modular protein immobilization based on the SpyCatcher/SpyTag chemistry.

12

Immobilization of SpyTagged proteins to SpyCatcher-PHA particles: Confirmation and

13

optimization of ligation reactions towards single and multi-protein display. To assess the

14

accessibility of the SpyCatcher domain displayed on PHA particles for ligation, i.e.,

15

immobilization of soluble free SpyTagged proteins via a spontaneous formation of a covalent

16

isopeptide linkage, we designed and produced several SpyTagged proteins, representing diverse

17

functionalities (Figure 1B). The selected proteins were the green fluorescent protein (GFP), a

18

biomarker commonly used in drug screening and diagnostic assays, the organophosphohydrolase

19

(OpdA), an organophosphate pesticide-degrading enzyme considered for bioremediation and the

20

Bacillus licheniformis α-amylase (BLA), a thermophilic industrially used starch-degrading

21

enzyme. BLA and GFP are monomeric, while OpdA needs to form a dimer to become active.

22

Purification of SpyTagged proteins for in vitro conjugation with the SP-P via SpyCatcher/SpyTag

23

chemistry was achieved by fusing a hexahistidine tag to the terminus suggested by the literature in


14

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Biomacromolecules

1

order to retain functionality of the respective protein and enable purification by subjecting the

2

tagged protein to Ni2+-NTA metal affinity chromatography.58-60 To avoid steric interference

3

between the SpyTag peptide and hexahistidine tag as well as retain the accessibility of both peptide

4

tags to their corresponding docking domains, we placed the peptide tags at opposite termini.

5

Therefore, in the current study, the SpyTag peptide was fused to the N-terminus for each soluble

6

protein. The yield, purity, apparent molecular weight and the identity of the SpyTagged proteins

7

were confirmed by SDS-PAGE (Figure 1E) and LC-MS/MS (Supporting Information Table S4).

8

Prior to protein ligation optimization, the SP fusion protein on PHA particles and all soluble

9

SpyTagged proteins were quantified by densitometry using a bovine serum albumin (BSA)

10

standard curve (Supporting Information Figures S1-S4). A linear curve could describe the BSA

11

standard curves generated for each densitometry analysis with r2 values of at least 0.98. Varying

12

dilution factors were used for each sample to ensure the readings are within the standard curve

13

linear range.

14

The various SpyTagged proteins were mixed with the SP-P as described in the Experimental

15

Section. SDS-PAGE analysis showed that after ligation an additional single protein band in lanes

16

2 to 4 with an apparent molecular weight greater than only SP fusion protein (68.4 kDa) appeared,

17

120.9 kDa, 104.0 kDa and 94.2 kDa for BLA-SP ligated protein (BLA-SP-L), OpdA-SP ligated

18

protein (OpdA-SP-L), and GFP-SP ligated protein (GFP-SP-L) respectively (Figure 1F). This step

19

also resulted in the production of single-protein immobilized SP-Ps: GFP immobilized SP-P (GFP-

20

SP-P), OpdA immobilized SP-P (OpdA-SP-P), and BLA immobilized SP-P (BLA-SP-P),

21

respectively. All ligation products were confirmed by tryptic peptide fingerprinting using LC/MS-

22

MS (Supporting Information Table S4). This result suggested successful immobilization of

23

SpyTagged proteins via ligation with SP-P. Hence, the SP-P provides a useful generic tool to


15


Page 16 of 52

1

immobilize different SpyTagged proteins. To further explore the tunability of the SP-P, we further

2

optimized the ligation reaction by varying the SpyCatcher-over-SpyTag reactant ratio and the

3

reaction time as shown in the supplementary material (Supporting Information Figures S5-S10).

4

Our optimization results showed that the ligation efficiency of SpyTag-to-SpyCatcher, i.e., the

5

percentage of total SpyTag-bearing proteins successfully ligated to the SpyCatcher domains on

6

PHA particles in the reaction mixture, of up to 83.2% could be achieved. The surface coverage of

7

SpyTagged proteins on SP-P varied from 19.0% to 59.0%.

8

To demonstrate the proof-of-concept that the various SpyTagged proteins can be immobilized

9

to the same SP-P, we implemented a step-by-step immobilization strategy as shown in the

10

schematic diagram provided in the supplementary material (Supporting Information Figure S11).

11

The subsequent successful ligation with each ligation step was monitored by SDS-PAGE analysis

12

of PHA particle associated proteins (Figure 1G). The gradual decrease in band intensity of SP

13

fusion protein (68.4kDa) correlated with the increasing formation of extra protein bands at the

14

higher molecular weight (120.9 kDa, 104.0 kDa & 94.2 kDa) representing the various ligation

15

products. These changes indicated that the step-wise loading of various soluble free SpyTagged

16

proteins onto the surface of SP-P reduced the availability of anchoring sites. The final ligation step

17

sample as shown in lane 4 of Figure 1G, where the SP fusion proteins were ligated with three

18

different SpyTagged proteins on the same PHA particle will be referred to as multifunctional SP-

19

P (MF-SP-P). We also attempted another strategy to prepare the MF-SP-P, where we incubated

20

the SP-P with equimolar quantities of SpyTagged proteins, however, this was less efficient

21

presumable due to undesirable steric competition between these SpyTagged proteins at anchoring

22

sites on SP-P. The protein surface coverage of SpyTagged OpdA immobilized on MF-SP-P was

23

greater than for SpyTagged GFP and SpyTagged BLA (Figure 1H). Since OpdA is a dimer, the


16

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Biomacromolecules

1

first ligated SpyTagged OpdA could sequester the second monomer via protein-protein interaction

2

and thereby facilitating ligation of SpyTagged OpdA onto MF-SP-P compared to other monomeric

3

proteins. A

SpyTag

SpyCatcher

Asp

Lys

rapid ligation

SpyCatchercoated PHA particles

Production and in vivo assembly of PHA particles in E. coli

D

C

B SIZE

SP

68.4 kDa

PS

69.1 kDa

PhaC

WT

55.5 kDa

PhaC

SpyTag – GFP – His6

25.8 kDa

SpyCatcher

M

kDa 235 170 130 93 70

SCHEMATIC DIAGRAM

FUSION PROTEIN

Functionalized SpyCatcherembedded PHA particles

In vitro mixing with SpyTagged proteins

PhaC SpyCatcher

1

2

3

Sample

mg nmol SpyCatcher SpyCatcher g wet per per particle g wet g wet per L particle particle

g cell mass per L

53

93 70

GFP

SpyTag – OpdA – His6

35.6 kDa

SpyTag

SpyTag – BLA – His6

52.5 kDa

SpyTag

H6

OpdA

H6

BLA

M

1

2

3

13.04

194.6

1.554

6.250

30

PS-P

2.79

41.9

1.681

8.619

22 18 14

WT-P

0

-

3.421

8.853

1

2

9

H6

F

E kDa 235 170 130

SpyTag

SP-P

41

G

kDa 235 170 130

M

1

2

3

4

H

kDa

M

1

2

3

4

kDa

235 170 130

93

70

70

93

53 70

53

41

41 41

53

30

9

30 22

Single protein immobilization SP only + BLA-SP-L GFP-SP-L

4

3

235 170 130 93

53

22 18 14

M

OpdA-SP-L

+

+

+

+

SP only BLA-SP-L

+

GFP-SP-L +

18 14

Step-by-step co-immobilization +

+

+

+

68.4 kDa

+

+

+

120.9 kDa

+

+

94.2 kDa

+

104.0 kDa

OpdA-SP-L

9

5

Figure 1. Design, production and modular functionalization of SpyCatcher-PhaC PHA

6

particles. (A) Schematic overview of production and modular functionalization of a generic

7

modular PHA platform for protein immobilization using SpyCatcher/SpyTag chemistry. (B)

8

Hybrid genes designed and used for this study. (C) SDS-PAGE analysis of SpyCatcher fusion


17


Page 18 of 52

1

protein displayed on PHA particles. Lane M, Gangnam pre-stained protein ladder; Lane 1, SP

2

fusion protein (68.4 kDa); Lane 2, PS fusion protein (69.1 kDa); Lane 3, WT protein (55.5 kDa).

3

(D) Production yields of SpyCatcher protein displayed on PHA particles. (E) SDS-PAGE analysis

4

of purified SpyTagged proteins. Lane M, Gangnam pre-stained protein ladder; Lane 1, SpyTagged

5

GFP (25.8 kDa); Lane 2, SpyTagged OpdA (35.6 kDa); Lane 3, SpyTagged BLA (52.5 kDa). (F)

6

SDS-PAGE analysis of various SpyTagged proteins immobilized on SP-P. Lane M, Gangnam pre-

7

stained protein ladder; Lane 1, SP fusion protein only; Lane 2, BLA-SP-L (120.9 kDa) & SP fusion

8

protein (68.4 kDa); Lane 3, GFP-SP-L (94.2 kDa) & SP fusion protein (68.4 kDa); Lane 4, OpdA-

9

SP-L (104.0 kDa) & SP fusion protein (68.4 kDa). (G) Visualization of step-by-step construction

10

of MF-SP-P by SDS-PAGE analysis. Lane M, Gangnam pre-stained protein ladder; Lane 1, SP

11

fusion protein only; Lane 2, BLA-SP-L (120.9 kDa) & SP fusion protein (68.4 kDa); Lane 3, BLA-

12

SP-L (120.9), GFP-SP-L (104.0 kDa) & SP fusion protein (68.4 kDa); Lane 4, BLA-SP-L (120.9

13

kDa, 20.1%), OpdA-SP-L (104.0 kDa, 24.3%), GFP-SP-L (94.2 kDa, 20.7%) & SP fusion protein

14

(68.4 kDa, 35.5%). (H) Comparison of different preparation strategies of MF-SP-P. Lane M,

15

Gangnam pre-stained protein ladder; Lane 1, SP only; Lane 2, Ligated proteins on MF-SP-P

16

prepared using equimolar quantities of SpyTagged proteins self-assembling strategy; Lane 3,

17

Ligated proteins on MF-SP-P prepared using step-wise reactant ratio modulated self-assembling

18

strategy; Purple arrow, BLA-SP-L; Red arrow, OpdA-SP-L; Green arrow, GFP-SP-L.

19 20

Furthermore, we analyzed the composition of the isolated SP-P against a pure poly (R)-3-

21

hydroxybutyric acid (PHB) standard using gas chromatography-mass spectrometry (GC/MS) as

22

shown in Figure 2A. This confirmed that PHB was produced. It also showed that the content of

23

PHB in SP-P was reduced by 28% when compared to the wild-type PHA particles (WT-P). This

24

implies a higher content of displayed protein over PHA mass upon fusion of the SpyCatcher

25

protein to the N-terminus of PhaC. This aligns with an increased protein band intensity of the SP

26

fusion protein as observed in Figure 1C when compared to that of the WT.


18

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Biomacromolecules

1

Moreover, we measured the Zeta potential of the SP-P, WT-P, and the functionalized SP-Ps

2

using electrophoretic light scattering (ELS) coupled with phase analysis light scattering (PALS)

3

(Figure 2B). We noticed the reduction of Zeta potential of the PHA particles at pH 7.5, from -16.9

4

± 0.6 mV to -29.3 ± 0.2 mV (mean ± 1 standard deviation (SD), n = 3) upon genetic fusion of

5

SpyCatcher domain to the N-terminus of PhaC when compared to the WT-P. Meanwhile, as was

6

expected, both SpyTagged GFP (-6.5 ± 1.7 mV) and SpyTagged BLA (-4.2 ± 0.8 mV) have net

7

negative Zeta potential values, and SpyTagged OpdA has a positive Zeta potential value (4.9 ± 1.1

8

mV) at pH 7.5 (mean ± 1SD, n = 3), where these proteins have an estimated isoelectric point of

9

6.26, 6.25 and 8.54, respectively.61 This result could further explain the faster ligation of

10

SpyTagged OpdA onto SP-P, compared to the others. Interestingly, we also noticed that the

11

immobilization of SpyTagged proteins onto the surface of SP-P via SpyCatcher/SpyTag chemistry

12

has no significant impact on the surface charge of SP-P. The Zeta potential of GFP-SP-P, OpdA-

13

SP-P, BLA-SP-P and MF-SP-P were -29.9 ± 1.2 mV, -31.5 ± 0.2 mV, -30.8 ± 0.3 mV and -30.2 ±

14

0.7 mV (mean ± 1SD, n = 3), respectively (Figure 2B).

15

We performed dynamic light scattering (DLS) analysis to determine the particle size and size

16

distribution of SP-P, WT-P and various functionalized SP-Ps (Figures 2C and 2D). Additionally,

17

particle distribution statistics are provided in the supplementary material (Supporting Information

18

Table S4). Statistically, SP-P has a larger Sauter mean diameter (D [3,2]) of 233 nm, and a lower

19

specific surface area of 24480 m2/kg, compared to those of WT-P. This discrepancy was mostly

20

due to the high polydispersity of the SP-P as shown in the particle size distribution (Figure 2C),

21

where SP-P tends to aggregate into two major aggregate clusters (about 1 μm and 10-20 μm),

22

which statistically increases the size of the SP-P. The undesirable formation of these aggregates

23

was presumably due to unspecific intermolecular and hydrophobic interactions, and likely


19


Page 20 of 52

1

independent of surface charges (Zeta potential), which were consistent across various

2

functionalized SP-Ps. It is also noteworthy to mention that the particle size distribution (Figure 2C)

3

suggested the individual SP-Ps have a smaller particle diameter (155 nm, blue arrow) when

4

compared to WT-P (259 nm, black arrow). Based on the DLS analysis, the amount of the

5

SpyCatcher domain displayed on SP-P was found to be approximately 0.091 fg per wet PHA

6

particle. Also, the SpyCatcher protein density at the surface of SP-P can be as close as around 8.4

7

× 1014 SpyCatcher domains per pm2.

8

Surprisingly, we found out that the SpyTagged protein-functionalized PHA particles (GFP-SP-

9

P, OpdA-SP-P, BLA-SP-P and MF-SP-P) are consistently more dispersed than those prior to

10

ligation, i.e., plain SP-P (Figure 2D). The D [3,2] of all functionalized SP-Ps ranged from

11

approximately 100 nm to 130 nm with the specific surface area ranging between about 41830

12

m2/kg to 54550 m2/kg, which was statistically significant increase compared to SP-P and WT-P

13

(Supporting Information Table S4). The large aggregate clusters of around 10-20 μm found in the

14

SP-P were strongly diminished for all functionalized SP-Ps and only low levels of aggregation

15

remained as shown in the particle size distribution, which suggested the surface functionalization

16

of SP-Ps reduced non-specific interactions between SP-Ps. As expected, we also noticed that

17

functionalization of SP-P with different SpyTagged proteins consistently increased the diameter

18

of individual SP-Ps from 155 nm (sky blue arrow) (Figure 2C) to approximately 180 nm to 200

19

nm (brown arrow) (Figure 2D) suggesting successful coating of SpyTagged proteins onto the SP-

20

Ps via the SpyCatcher/SpyTag chemistry without affecting the assembled structure of the SP-Ps.


20

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Biomacromolecules

1 2

Figure 2. Physicochemical characterization of SpyCatcher-PhaC PHA particles. (A)

3

Compositional analysis of PHA particles by GC/MS. (B) Zeta potential of PHA particles and

4

soluble SpyTagged proteins by ELS/PALS (mean ± 1SD, n = 3). (C) Particle size distribution of

5

SP-P and WT-P (mean ± 1SD, n = 3). (D) Particle size distribution of various functionalized SP-

6

Ps by DLS analysis (mean ± 1SD, n = 3).

7 8

Tunable protein immobilization using the SP-P platform was achieved by varying the

9

SpyCatcher:SpyTag ratios, which enabled control of the amount of SpyTagged proteins ligated to


21


Page 22 of 52

1

SP-P. However, ligation reactions using increasing amounts of SpyTagged protein over

2

SpyCatcher indicated that a significant fraction of SP fusion proteins remained unligated. Thus,

3

the current conditions did not result in complete saturation of SP-P with the SpyTagged proteins

4

of interest (Supporting Information Figures S5-S10). Possibly PHA particle aggregation might

5

have restricted accessibility of SpyTagged proteins to the SpyCatcher domain displayed on PHA

6

particles. Besides, the protein surface properties such as Zeta potential and hydrophobicity as well

7

as the accessibility of the SpyTag itself could have interfered with the ligation reaction. Notably,

8

the electrostatic attraction between the positively charged SpyTagged OpdA and the negatively

9

charged SP-P facilitated ligation between the SpyCatcher domain and SpyTag, when compared to

10

both negatively-charged SpyTagged GFP and BLA, which were also used in this study.

11

In fact, previous studies described these ligation efficiency issues, when immobilizing

12

SpyTagged proteins onto other SpyCatcher-supporting scaffolds. Thrane et al. noted that coupling

13

efficiency of several antigens onto the Spy-VLPs (SpyCatcher embedded virus-like particles)

14

ranged from 33% to 88% and suggested that small proteins are less likely to be affected by steric

15

hindrance during the ligation process.44 Meanwhile, Jia et al. also found a similar problem, where

16

higher amounts of SpyCatcher proteins are needed to enhance the conjugation efficiency.47 Apart

17

from raising the issue that large proteins are more susceptible to steric hindrance, they also argued

18

that it might be due to the SpyCatcher proteins are being trapped within the hyperbranched

19

structure of the SpyCatcher polymer, which in turn limited the accessibility of SpyTagged proteins

20

to interact with SpyCatcher protein.47

21

For initial validation of successful modular functionalization of SP-P, i.e., to assess whether the

22

functionality of the SpyTagged proteins can be retained after immobilization on PHA particles, we

23

first screened for fluorescence of immobilized GFP on the SP-P using the soluble free GFP as


22

Page 23 of 52 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

1

positive control as shown in Figure 3. To determine the amount of GFP immobilized on the PHA

2

particles, both GFP-SP-P, and MF-SP-P were subjected to SDS-PAGE analysis followed by

3

densitometry analysis (Supporting Information Figures S12-S13). Both GFP-SP-P and MF-SP-P

4

in suspension emitted bright green fluorescence comparable to the soluble GFP prior to

5

sedimentation by centrifugation (Figure 3B (top)). On the contrary, we noticed that the negative

6

controls (WT-P and SP-P) did not emit the same intensity of fluorescence. Brighter fluorescence

7

can be seen visually upon localization of particles by physical means, i.e., centrifugation (Figure

8

3B (bottom)). Additional fluorescence screening images can be found in the supporting material

9

(Supporting Information Figures S16-S17). We measured the fluorescence intensity of the samples

10

as described in the Experimental Section. From the bar graph in Figure 3C, the fluorescence

11

intensity of both GFP-SP-P and MF-SP-P did show an equivalent signal compared to free GFP.

12

At the microscopic level, as shown in Figure 3D, both GFP-coated PHA particles also exhibited

13

high local fluorescence on the PHA particles. These results indicate successful modular

14

functionalization of SP-P using SpyTagged GFP.


23

Biomacromolecules

B

C

EXCITATION

Fluorescence Arbitrary Units

A

CENTRIFUGATION

4500 4000 3500 3000 2500 2000 1500 1000 500 0

FLUORESCENCE

D SP-P (negative)

GFP (positive)

Controls

WT-P (negative)

100#μm

100#μm

100#μm

MF-SP-P

GFP-SP-P

Immobilized GFP

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 52

100#μm

100#μm

1 2

Figure 3. Fluorescence of GFP immobilized to SpyCatcher-PhaC PHA particles. (A)

3

Schematic of immobilized GFP on the SP-P upon exposure to excitation light. (B) Fluorescence

4

can be detected on immobilized GFP anchored on SP-P. (C) Arbitrary fluorescence intensity of

5

the GFP-SP-P and MF-SP-P with controls (mean ± 1 SD, n = 3). (D) Fluorescence microscopy

6

analysis of the GFP-SP-P and MF-SP-P with controls.

7


24

Page 25 of 52 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

1

Enzyme immobilization using SpyCatcher-PHA particles. We tested the utility of SP-P as

2

supporting material for one or two enzymes. We choose OpdA and BLA for this study because of

3

their vast potential in industrial and agricultural applications. SpyTagged enzymes were produced,

4

purified and immobilized at SpyCatcher:SpyTag ratios of 4:1 and 3:1 for OpdA and BLA,

5

respectively, to achieve an approximately 20% surface coverage on SP-P, which give us OpdA-

6

SP-P, BLA-SP-P and MF-SP-P. The SDS-PAGE analysis confirmed the successful

7

immobilization of these enzymes on SP-P and densitometry was used to quantify enzymes

8

immobilized to PHA particles (Supporting Information Figures S13-S15). The functionality of

9

immobilized and free BLA was first qualitatively assessed using 1% (wt/vol) starch agar

10

(Supporting Information Figure S18). All BLA containing samples created a clear transparent

11

zone, which indicated starch hydrolysis. Then, we tested the enzyme activities of both immobilized

12

and soluble free forms in their respective reaction mixture. We compared the substrate conversion

13

rates of immobilized enzymes to those of purified soluble enzymes (Figure 4). Our findings

14

suggested that both immobilized OpdA and BLA outperformed their soluble counterparts. The

15

catalytic activity of immobilized OpdA on OpdA-SP-P (5.09 ± 0.08 U/mg) was around 9% higher

16

when compared to free OpdA (4.66 ± 0.26 U/mg). Interestingly, we found that the immobilized

17

OpdA on MF-SP-P (6.33 ± 0.16 U/mg) exhibited a much faster coumaphos degradation when

18

compared to both OpdA-SP-P and soluble OpdA (mean ± 1SD, n = 3). Meanwhile, the specific

19

BLA activities of immobilized BLA on BLA-SP-P and MF-SP-P were 3.72 ± 0.03 U/mg and 3.67

20

± 0.04 U/mg, respectively, which was approximately 30% higher than the soluble BLA activity of

21

2.63 ± 0.07 U/mg (mean ± 1SD, n = 3).

22

These results suggested that the spatial organization and oriented display of enzymes on SP-P

23

can increase the rate of the catalytic reaction. Also, the close-proximity between immobilized


25


Page 26 of 52

1

enzymes on the SP-P might have created macromolecular crowding effects, leading to an enhanced

2

substrate conversion rate of the enzymes studied. Kao et al. made a similar observation and showed

3

that clustering of immobilized lysozyme on mesoporous silica nanoparticles results, due to

4

artificially created crowded microenvironment, a better performance than the soluble

5

counterpart.62 Yang et al. also reported that the catalytic efficiency of both 7α-hydroxysteroid

6

dehydrogenase and 7β-hydroxysteroid dehydrogenase could be increased by controlling the

7

crowding density of immobilized enzymes of interest on chitosan-epoxy resin carriers.9 However,

8

it may vary from case to case, as different proteins have different characteristics and surface

9

clustering of some proteins on supporting scaffolds might have an adverse effect due to topological

10

frustration and steric hindrance.63-65


26

Page 27 of 52

B

A

250

[Chlorferon] (μM)

Substrate Coumaphos

Products Chlorferon (Fluorescence↑)

200 150 WT-P SP-P OpdA OpdA-SP-P MF-SP-P

100 50 0

DETP

0

Immobilized OpdA

20

40

60 80 Time (min)

100

120

D

C Immobilized BLA

350 300 [Nitrophenol] (μM)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

250 200 WT-P SP-P BLA BLA-SP-P MF-SP-P

150 100 50

Substrate Ethylidene-pNP-G7

1

Product Nitrophenol (OD405 nm)

0 0

30

60

90 120 Time (min)

150

180

2

Figure 4. OpdA and BLA enzymatic functionality assays. (A) Schematic illustration of OpdA

3

activity assay using coumaphos as substrate. Coumaphos is degraded into chlorferon and

4

dietlythiophosphate (DETP) by immobilized OpdA. (B) Reaction time course of coumaphos

5

degradation hydrolyzed by OpdA-SP-P and MF-SP-P with appropriate controls (mean ± 1SD, n =

6

3). (C) Schematic illustration of BLA activity determined by 3,5-dinitrosalicyclic acid colorimetric

7

assay. Nitrophenol is released from ethylidene-pNP-G7 by enzymatic activity of immobilized

8

BLA. (D) Reaction time course of ethylidene-pNP-G7 degradation hydrolyzed by BLA-SP-P and

9

MF-SP-P with appropriate controls (mean ± 1SD, n = 3).

10


27


Page 28 of 52

1

Alteration of material surface microenvironments upon covalent docking of enzymes could

2

significantly affect the performance of immobilized enzymes, which might explain enhanced

3

OpdA performance of MF-SP-P. A combination of surface charge, hydrophobicity/hydrophilicity,

4

surface topology and orientation of active sites among enzymes and/or the material-protein

5

interface allow the construction of favorable microenvironments, which could contribute to the

6

enhanced performance and stability of an enzyme.5 The decoration of different functional proteins

7

on MF-SP-P might have created a favorable microenvironment to degrade coumaphos using

8

OpdA, in such that coumaphos is channeled onto the active sites of immobilized OpdA on MF-

9

SP-P. This phenomenon might explain the improved coumaphos conversion rate of MF-SP-P over

10

OpdA-SP-P in degrading coumaphos, unlike the case of immobilized BLA on BLA-SP-P and MF-

11

SP-P where no significant difference in substrate degradation rate was measured.

12

The substrate conversion rate of OpdA ligated to SP-P outperformed the OpdA displayed on

13

PHA particles via direct translational fusion with PhaC. Blatchford et al. reported up to 23%

14

reduction in coumaphos conversion rate of PhaC-OpdA beads when compared to free OpdA.66

15

BLA displayed via translational fusion with PhaC on BLA-PhaC beads retained the original

16

substrate conversion rate of soluble BLA.54 However, Figure 4 shows the enhanced performance

17

at varying rates for both immobilized enzymes ligated to SP-P when compared with their soluble

18

counterparts. As mentioned, implementation of this modular approach for immobilization of

19

enzymes could eliminate potential protein misfolding and orientation issues that often happen with

20

surface-displayed proteins on various support materials.28-29, 67-68 Also, the observed increase in

21

catalytic performance of immobilized enzymes using the modular system as described in this study

22

versus the PhaC fusion based approach could be due to the changes in physiochemical properties

23

such as reduced particle size, which in turn led to a larger surface area over volume ratio of the


28

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Biomacromolecules

1

particles. Fusion of SpyCatcher protein to the N-terminus of PhaC reduced the particle size of the

2

individual PHA particles (155 nm) when compared to the wild-type PHA particles (259 nm) as

3

mentioned above. Rubio-Reyes et al. noticed the variation in particle size, ranging from 500 nm

4

to 750 nm when different antigens were displayed on PHA beads.24 González-Miro et al. also

5

reported that the particle size of PHA inclusions decreased from 500 nm to 100 nm upon fusion of

6

PsaA to the PhaC.30 These observations indicate that fusing different proteins to PhaC impacts on

7

the size and size distribution of PHA particles. However, functionalization of SP-P with various

8

SpyTagged proteins via SpyCatcher/SpyTag chemistry has minimal impact on the particle size on

9

top of the remarkable consistency in the particle dispersity as demonstrated earlier. Therefore, this

10

genetic fusion partner dependent variability can be reduced by exploiting the modular SpyCatcher-

11

PHA particle approach towards the development of a generic protein-immobilizing platform.

12

Thermal stability. We evaluated the thermal stability of the immobilized and free SpyTagged

13

proteins for their functional performance at varying pre-incubation temperatures (Figures 5A-5C),

14

as described in the Experimental Section. The modular immobilization of SpyTagged proteins

15

retained the inherent thermal stability of the soluble form. We observed the same loss of

16

fluorescence intensity in both immobilized and soluble free GFP as shown in Figure 5A. Rapid

17

loss of fluorescence in all samples started at 75°C and was abolished at 85°C, which is in good

18

agreement with the reported values of the GFP melting temperature (76°C - 78°C).69-70 This

19

observation indicates that the immobilized GFP retained the thermal stability of free GFP.

20

Moreover, immobilized OpdA retained the thermal stability of soluble free OpdA, as illustrated

21

in Figure 5B. In general, the substrate conversion rate of immobilized and free OpdA remained

22

stable until 65°C, consistent with the reported apparent melting point of free OpdA and PhaC-

23

OpdA PHA beads.66 Interestingly, we detected an early decline in OpdA performance on the MF-


29


Page 30 of 52

1

SP-P, where we observed about 22% loss in enzymatic performance as a result of 10°C rise in

2

temperature from 45°C to 55°C. This observation can be explained by the thermal dissociation of

3

unique surface microenvironment properties created on the MF-SP-P mentioned above.

4

Presumably, loss of the advantageous surface properties that facilitate better coumaphos

5

degradation resulted in a reduced catalytic performance of MF-SP-P, back to the levels similar to

6

OpdA-SP-P.

7

We noted an increased substrate conversion rate of free BLA at temperatures ranging from 55°C

8

to 85°C as shown in Figure 5C, in line with several studies reported.71-73 Overall, immobilized

9

BLA on both BLA-SP-P and MF-SP-P showed the same thermal stability as soluble BLA.

10

However, immobilized BLA on MF-SP-P appeared to be slightly more susceptible to thermal

11

degradation in the high-temperature range, when compared with BLA-SP-P and soluble BLA. This

12

phenomenon could be due to the structural destabilization and shielding of active sites caused by

13

the other immobilized proteins unfolding at elevated temperatures. Although BLA-SP-P and

14

soluble BLA were still relatively stable at 85°C, we noticed an approximate 14% reduction in

15

performance of BLA on MF-SP-P. We also found that a further increase in temperature beyond

16

85°C, to 95°C, resulted in around 27%, 16% and 35% loss of degradation performance for soluble

17

BLA, BLA-SP-P, and MF-SP-P, respectively. Interestingly, immobilized BLA in both BLA-SP-P

18

and MF-SP-P maintained high levels of performance in the low-temperature range, while higher

19

temperatures were required for BLA immobilized by direct translational fusion to PhaC on PHA

20

beads. The performance of BLA-PhaC beads produced by Rasiah and Rehm showed similar

21

temperature dependency to that of the soluble free BLA.54 This finding suggested that a broader

22

optimum working temperature range for thermostable enzymes can be achieved by using the

23

SpyCatcher-PHA particle approach.


30

Page 31 of 52

A

C

B 250

250

200

200

3000 2500

WT-P SP-P GFP GFP-SP-P MF-SP-P

2000 1500 1000

150 WT-P SP-P OpdA OpdA-SP-P MF-SP-P

100 50

500 0

[Nitrophenol] (μM)

3500 [Chlorferon] (μM)


4000

0

5 15 25 35 45 55 65 75 85 95 Temperature (oC)

WT-P SP-P GFP GFP-SP-P MF-SP-P

2000 1500 1000

5 15 25 35 45 55 65 75 85 95 Temperature (ºC)

250

200 150

[Nitrophenol] (μM)

[Chlorferon] (μM)


2500

50

15 25 35 45 55 65 75 85 95 Temperature (ºC)

WT-P SP-P OpdA OpdA-SP-P MF-SP-P

3500 3000

WT-P SP-P BLA BLA-SP-P MF-SP-P

100

F 250

4000

150

0 5

E

D

100 50

200 150

WT-P SP-P BLA-P BLA-SP-P MF-SP-P

100 50

500 0

0 3

5

7 pH

9

5

7 pH

9

11

3

350

250

300

2000 1500 1000 500

200 150 100 50

SP-P

2

3 Cycle

GFP-SP-P

4

5 MF-SP-P

7 pH

9

11

1 WT-P

250 200 150 100 50 0

0 1

WT-P

[Nitrophenol] (μM)

300

3000

[Chlorferon] (μM)

3500 2500

5

I

H

0

1

0 3

11

G Fluorescence Arbitrary Units

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

SP-P

2

3 Cycle OpdA-SP-P

4

1

5 MF-SP-P

WT-P

SP-P

2

3 Cycle BLA-SP-P

4

5 MF-SP-P

2

Figure 5. Stability and reusability of SpyTagged proteins immobilized to SpyCatcher-PhaC

3

PHA particles. (A) Arbitrary fluorescence intensity of immobilized GFP on GFP-SP-P and MF-

4

SP-P with controls at varying temperatures (mean ± 1SD, n = 3). (B) Amount of chlorferon

5

released from coumaphos hydrolyzed by immobilized OpdA on OpdA-SP-P and MF-SP-P with

6

controls at varying temperatures (mean ± 1SD, n = 3). (C) Amount of nitrophenol liberated from

7

ethylidene-pNP-G7 by immobilized BLA on BLA-SP-P and MF-SP-P with controls at varying

8

temperatures (mean ± 1SD, n = 3). (D) Arbitrary fluorescence intensity of immobilized GFP on

9

GFP-SP-P and MF-SP-P with controls at varying pH values (mean ± 1SD, n = 3). (E) Amount of

10

chlorferon released from coumaphos hydrolyzed by immobilized OpdA on OpdA-SP-P and MF-

11

SP-P with controls at varying pH values (mean ± 1SD, n = 3). (F) Amount of nitrophenol liberated

12

from ethylidene-pNP-G7 by immobilized BLA on BLA-SP-P and MF-SP-P with controls at


31


Page 32 of 52

1

varying pH values (mean ± 1SD, n = 3). (G) Arbitrary fluorescence intensity of immobilized GFP

2

on GFP-SP-P and MF-SP-P with controls over five cycles (mean ± 1SD, n = 3). (H) Amount of

3

chlorferon released from coumaphos hydrolyzed by immobilized OpdA on OpdA-SP-P and MF-

4

SP-P with controls over five cycles (mean ± 1SD, n = 3). (I) Amount of nitrophenol liberated from

5

ethylidene-pNP-G7 by immobilized BLA on BLA-SP-P and MF-SP-P with controls over five

6

cycles (mean ± 1SD, n = 3).

7 8

pH stability. The stability of enzymes in acidic and alkaline environments is of interest as it

9

impacts on their application in various bioprocesses. To determine the pH stability of both

10

immobilized and soluble proteins, we exposed the various proteins to different pH values ranging

11

from pH 3.0 to pH 11.0 and then assessed their functionality (Figures 5D-5F). GFP is known to be

12

relatively stable in weak alkaline solutions but degrades at acidic conditions, consistent with the

13

pH stability profile of free GFP as shown in Figure 5D.74 We observed a similar trend for

14

immobilized GFP on both GFP-SP-P and MF-SP-P over the pH range studied. However, we noted

15

up to around 17% reduction of the signal of immobilized GFP on MF-SP-P in alkaline conditions,

16

compensated with higher fluorescence at pH 3. Although the causes for this outcome are not

17

understood, we can interpret this observation as being a result of steric effects caused by the other

18

immobilized proteins, which lead to the minimal shift in endurance against pH-triggered

19

destabilization. Jin et al. reported a similar observation for co-immobilization of chloroperoxidase

20

(CPO) and horseradish peroxidase (HRP) on zinc oxide-silicon dioxide composite scaffolds, where

21

reduced enzymatic activity was observed for the co-immobilized peroxidases at their respective

22

optimum pH values of pH 3 and 6.75 However, this was compensated with higher performance at

23

pH 5 and better tolerance against pH fluctuations.75


32

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Biomacromolecules

1

As shown in Figure 5E, the SP-P stabilized the immobilized OpdA on both OpdA-SP-P and MF-

2

SP-P at alkaline conditions, as only about 27% and 32% loss in substrate conversion rate at pH 9

3

was observed when compared to the performance at optimum condition at neutral pH while soluble

4

OpdA showed approximately 55% reduction in substrate conversion rate. We observed a notable

5

further deterioration in the OpdA catalytic functionality at pH 11 for all the samples. Though half

6

of the maximum OpdA performance was retained for OpdA-SP-P and MF-SP-P at pH 11, only

7

around 20% of the optimum performance was retained for the soluble counterpart. Poor resistance

8

against low pH values of free OpdA in this study is consistent with previous findings.76 We found

9

improved stability at low pH stress for OpdA-SP-P and MF-SP-P at pH 5, where approximately

10

half of the functionality was still present, while only about 32% of the substrate conversion rate

11

was retained for soluble free OpdA. These data suggested that ligation of OpdA to SP-P strongly

12

increased pH stability. Venning-Slatter et al. also observed the strong loss of OpdA relative

13

catalytic performance at pH 3, who reported a diminished performance of OpdA functionality at

14

pH 3 with the OpdA-immobilized to PHA particles or GFP particles using translational fusions for

15

in situ immobilization. However, their claim that OpdA displayed on both PHA or GFP particles

16

were able to withstand a broader pH range was not found for OpdA immobilized to PHA particle

17

via ligation.77 Shorter pre-incubation time (10 min) of their samples at varying pH buffers could

18

explain this difference. We pre-treated our samples with different buffers for a longer time (30

19

min) before subjecting them to the functional assay. Our results are in good agreement with those

20

obtained by Milani et al and Tang et al, where they pre-incubated their immobilized OpdA samples

21

for 5 h and 1 h, respectively.78-79

22

It is also noteworthy to find that immobilized BLA on SP-P are less susceptible to pH

23

inactivation, especially BLA-SP-P as shown in Figure 5F. On the contrary, only around 30% of


33


Page 34 of 52

1

the optimum substrate conversion rate was retained for soluble free BLA at both low and high pH

2

levels, which was in agreement with previously published results.71, 73 The stability of BLA ligated

3

to SP-P resembled what had been achieved by using in situ immobilization of BLA to PHA

4

particles.54, 77 While BLA-SP-P and MF-SP-P retained most of the optimal catalytic performance

5

of BLA (approximately 92% and 72%, respectively) at pH 3, BLA-PHA particles performed

6

poorly at the same pH value.

7

Overall, the pH stability profile of the proteins immobilized to SP-P via ligation was improved,

8

particularly at higher pH values. Apart from the covalent bond formed between the SpyCatcher

9

domain and the SpyTag peptide, nonspecific interactions due to macromolecular crowding

10

between proteins80-81, and possibly with the scaffolding material could have a stabilizing effect.

11

Using on-surface circular dichroism spectroscopy, White et al. analytically demonstrated that the

12

macromolecular

13

(AQLKKKLQANKKKLAQLKWKLQALKKKLAQGGGSC) via covalent attachment onto

14

thiol-reactive surfaces, drastically shifted the threshold pH for the conformational change of

15

BASE-C from random-coil into α-helical structure (pH 9 in soluble free state to higher than pH 4

16

in immobilized state). They suggested that the densely packed BASE-C on the supporting scaffold,

17

which created the excluded volume effect, drives the change in protein folding via hydrophobic

18

interactions.82 In the case of immobilized GFP and OpdA, the nonspecific stabilizing effects such

19

as electrostatic and hydrophobic interactions might have been disrupted under acidic conditions.

20

Consequently, the stabilizing effects created might be significantly impaired, allowing the

21

ionization of crucial amino acid residues that constitute the active catalytic sites or are involved in

22

electrostatic interactions. However, the stabilizing mechanism for immobilized BLA could

23

withstand low pH conditions, retaining at least 80% of the BLA functionality.

crowding

of

a

synthetic

biomolecule,

BASE-C

peptide


34

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Biomacromolecules

1

Reusability. Although proteins are widely used for a variety of medical and industrial

2

applications, it can be challenging to use them in the continuous processing because of insufficient

3

stability as well as of difficulties in separating proteins from the bulk environment for reuse.

4

However, protein immobilization techniques make the use of enzymes adaptable to the current

5

continuous processing technologies by facilitating recovery and reuse.83-84 To demonstrate that

6

proteins ligated to SP-P are reusable, we repeated the functional assay of the respective samples

7

in five cycles. Figures 5G-5I show the comparison of the repeated use of the immobilized proteins.

8

Respective soluble proteins were recovered by ultrafiltration, and their reusability is shown in

9

Supporting Information Figure S11. Overall, the immobilized proteins showed a similar loss of

10

functionality when compared to the respective soluble proteins over five cycles. As shown in

11

Figure 5G, immobilized GFP on GFP-SP-P and MF-SP-P retained about 92% and 91% of the

12

initial fluorescence signal over the cycles, while the soluble counterpart retained approximately

13

91% (Supporting Information Figure S19).

14

Similarly, we observed a slightly improved reusability for OpdA. Figure 5H indicates a notable

15

increment in reusability of the immobilized form of this enzyme, where around 83% and 86% of

16

OpdA catalytic functionality was retained for OpdA-SP-P and MF-SP-P respectively, compared

17

to 80.3% of that in free OpdA (Supporting Information Figure S19). Venning-Slater et al. reported

18

that the OpdA-displaying GFP particles retained about 81% of the substrate conversion rate after

19

seven cycles, which is comparable to our findings.77 Our results suggested that OpdA ligated to

20

SP-P showed better reusability performance when compared to OpdA immobilized to polyamide

21

nanofibrous scaffold and glutaraldehyde-crosslinked chitosan beads where approximately 60% of

22

OpdA functionality was retained after five repeated uses.78, 85


35


Page 36 of 52

1

A slight loss of BLA catalytic performance was found for both BLA-SP-P and MF-SP-P, as about

2

73% and 69% of substrate conversion rate was retained (Figure 5I), respectively, while soluble

3

BLA retained around 76% of the native functionality (Supporting Information Figure S19).

4

However, our results are consistent with previous studies conducted by Gangadharan et al. and

5

Radovanović et al., where an approximately 30% loss in functionality over five cycles was

6

reported.86-87 Additionally, immobilized BLA exhibited a better reusability performance than

7

obtained with BLA displayed on self-assembled protein particles, where less than 10% of the

8

optimum performance was retained at the fourth cycle.77

9 10

CONCLUSIONS

11

In this study, we developed a versatile modular platform for protein immobilization by merging

12

the in vivo PHA particle display technology with the in vitro SpyCatcher/SpyTag chemistry.

13

SpyTagged functional proteins can be anchored onto the SpyCatcher-displaying PHA particles via

14

rapid formation of a covalent isopeptide bond by mixing the two components. Our results also

15

revealed that this modular platform shows versatility and tunability, by controlling the reactant

16

molar ratio when ligating different functional proteins with SpyCatcher-PHA particles, which

17

leads to convenient co-immobilization of multiple protein functions reconstituting multiprotein

18

complexes at the surface of PHA particles. This study demonstrated that multifunctionality could

19

be easily achieved using our SpyCatcher-PHA particles platform. Both macromolecular crowding

20

and creation of favorable microenvironments on the surface of the scaffolding material as well as

21

oriented display could explain the overall retained or enhanced functionality and stability. In

22

contrast to the in vivo PHA particle immobilization techniques, the SpyCatcher-PHA particle


36

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Biomacromolecules

1

approach reduces the risk of misfolding due to separate production and by using the most suitable

2

production host for the target protein and the PHA support material. The SpyCatcher-PHA particle

3

approach offers a generic protein immobilization platform where SpyTagged target proteins can

4

be efficiently ligated to a polymeric support material without the need of costly chemical cross-

5

linkers or enzymes.

6 7 8 9 10 11 12 13 14 15 16 17 18


37


Page 38 of 52

1

ASSOCIATED CONTENT

2

Supporting Information.

3

The following files are available free of charge.

4

Supporting info - Wong and Rehm, 2018.pdf: Bacterial strains, plasmids, primers used in this

5

study; Plasmid construction strategy; Identification of fusion proteins by liquid chromatography-

6

tandem mass spectrometry (LC-MS/MS); Densitometric protein quantification of SP fusion

7

protein on PHA particles and SpyTagged proteins for SpyCatcher/SpyTag chemistry ligation

8

optimization; Ligation optimization of SpyTagged proteins onto SpyCatcher-PhaC PHA particles;

9

Schematic overview of stepwise multifunctionalization of SpyCatcher-PhaC PHA particles;

10

Particle size distribution statistics of various PHA particles used in this study; Densitometric

11

protein quantification of immobilized SpyTagged proteins on various functionalized SpyCatcher-

12

PhaC PHA particles. Additional images of fluorescence screening of immobilized GFP on

13

SpyCatcher-PhaC PHA particles; Starch degradation screening of the immobilized BLA on

14

SpyCatcher-PhaC PHA particles; Recyclability of soluble free SpyTagged proteins as positive

15

control.

16

AUTHOR INFORMATION

17

Corresponding Author

18

*Bernd H. A. Rehm. Email: [email protected]. Phone: +61737354233

19

Author Contributions

20

The manuscript was written through contributions of all authors. All authors have given approval

21

to the final version of the manuscript.


38

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Biomacromolecules

1

Funding Sources

2

This work was supported by Griffith University (Australia), the MacDiarmid Institute of Advanced

3

Materials and Nanotechnology (New Zealand), and Institute of Fundamental Sciences, Massey

4

University (New Zealand).

5

ACKNOWLEDGMENT

6

The authors would like to thank the financial support from MacDiarmid Institute of Advanced

7

Materials and Nanotechnology (New Zealand) and Institute of Fundamental Sciences, Massey

8

University (New Zealand) and Griffith University (Australia). The authors are also thankful to

9

Trevor Loo for his assistance with LC-MS/MS analysis.

10

ABBREVIATIONS

11

PHA, Polyhydroxyalkanoates; PhaC, PHA synthase; SP, SpyCatcher-PhaC fusion protein; PS,

12

PhaC-SpyCatcher fusion protein; WT, wild-type PhaC; SP-P, SpyCatcher-PhaC PHA particles;

13

PS-P, PhaC-SpyCatcher PHA particles; WT-P, wild-type PhaC PHA particles; GFP, green

14

fluorescent protein; OpdA, organophosphohydrolase; BLA, Bacillus licheniformis α-amylase;

15

GFP-SP-L, GFP and SpyCatcher-PhaC ligated protein; OpdA-SP-L, OpdA and SpyCatcher-PhaC

16

ligated protein; BLA-SP-L, BLA and SpyCatcher-PhaC ligated protein; GFP-SP-P, GFP

17

immobilized SpyCatcher-PhaC PHA particles; OpdA-SP-P, OpdA immobilized SpyCatcher-PhaC

18

PHA particles; BLA-SP-P, BLA immobilized SpyCatcher-PhaC PHA particles; MF-SP-P,

19

multifunctional SpyCatcher-PhaC PHA particles.

20 21 22


39


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REFERENCES (1) Hoarau, M.; Badieyan, S.; Marsh, E. N. G. Immobilized enzymes: understanding enzyme– surface interactions at the molecular level. Organic & Biomolecular Chemistry 2017, 15 (45), 9539-9551. (2) Rodrigues, R. C.; Ortiz, C.; Berenguer-Murcia, Á.; Torres, R.; Fernández-Lafuente, R. Modifying enzyme activity and selectivity by immobilization. Chemical Society Reviews 2013, 42 (15), 6290-6307. (3) Zhao, Z.; Fu, J.; Dhakal, S.; Johnson-Buck, A.; Liu, M.; Zhang, T.; Woodbury, N. W.; Liu, Y.; Walter, N. G.; Yan, H. Nanocaged enzymes with enhanced catalytic activity and increased stability against protease digestion. Nature Communications 2016, 7, 10619. (4) Fu, J.; Liu, M.; Liu, Y.; Woodbury, N. W.; Yan, H. Interenzyme substrate diffusion for an enzyme cascade organized on spatially addressable DNA nanostructures. Journal of the American Chemical Society 2012, 134 (12), 5516-5519. (5) Talbert, J. N.; Goddard, J. M. Enzymes on material surfaces. Colloids and Surfaces B: Biointerfaces 2012, 93, 8-19. (6) Zhang, D.-H.; Yuwen, L.-X.; Peng, L.-J. Parameters affecting the performance of immobilized enzyme. Journal of Chemistry 2013, 2013. (7) Guzik, U.; Hupert-Kocurek, K.; Wojcieszyńska, D. Immobilization as a strategy for improving enzyme properties-application to oxidoreductases. Molecules 2014, 19 (7), 89959018. (8) Schroeder, M. M.; Wang, Q.; Badieyan, S.; Chen, Z.; Marsh, E. N. G. Effect of surface crowding and surface hydrophilicity on the activity, stability and molecular orientation of a covalently tethered enzyme. Langmuir 2017, 33 (28), 7152-7159. (9) Yang, Q.; Wang, B.; Zhang, Z.; Lou, D.; Tan, J.; Zhu, L. The effects of macromolecular crowding and surface charge on the properties of an immobilized enzyme: activity, thermal stability, catalytic efficiency and reusability. RSC Advances 2017, 7 (60), 38028-38036. (10) Balcells, C.; Pastor, I.; Vilaseca, E.; Madurga, S.; Cascante, M.; Mas, F. Macromolecular crowding effect upon in vitro enzyme kinetics: mixed activation–diffusion control of the oxidation of NADH by pyruvate catalyzed by lactate dehydrogenase. The Journal of Physical Chemistry B 2014, 118 (15), 4062-4068. (11) Zaak, H.; Siar, E.-H.; Kornecki, J. F.; Fernandez-Lopez, L.; Pedrero, S. G.; Virgen-Ortíz, J. J.; Fernandez-Lafuente, R. Effect of immobilization rate and enzyme crowding on enzyme stability under different conditions. The case of lipase from Thermomyces lanuginosus immobilized on octyl agarose beads. Process Biochemistry 2017, 56, 117-123. (12) Morán-Zorzano, M. T.; Viale, A. M.; Muñoz, F. J.; Alonso-Casajús, N.; Eydallín, G. G.; Zugasti, B.; Baroja-Fernández, E.; Pozueta-Romero, J. Escherichia coli AspP activity is enhanced by macromolecular crowding and by both glucose‐1, 6‐bisphosphate and nucleotide‐sugars. FEBS Letters 2007, 581 (5), 1035-1040. (13) Jia, J.; Peng, X.; Qi, W.; Su, R.; He, Z. Effects of macromolecular crowding on alkaline phosphatase unfolding, conformation and stability. International Journal of Biological Macromolecules 2017, 101, 373-382. (14) Minton, A. P. Effect of a concentrated “inert” macromolecular cosolute on the stability of a globular protein with respect to denaturation by heat and by chaotropes: a statisticalthermodynamic model. Biophysical Journal 2000, 78 (1), 101-109.


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Design of Modular Polyhydroxyalkanoate Scaffolds for Protein

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