Immobilization of Recombinant E. coli Cells in a Bacterial Cellulose

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Immobilization of Recombinant E. coli cells in a Bacterial Cellulose-Silk Composite Matrix to Preserve Biological Function Irina Drachuk, Svetlana V Harbaugh, Ren D. Geryak, David L Kaplan, Vladimir V. Tsukruk, and Nancy Kelley-Loughnane ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00367 • Publication Date (Web): 23 Aug 2017 Downloaded from http://pubs.acs.org on August 24, 2017

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Immobilization of Recombinant E. coli cells in a Bacterial Cellulose-Silk Composite Matrix to Preserve Biological Function Irina Drachuk*,†,‖‖ Svetlana Harbaugh,‡, ‖‖ Ren Geryak,§ David L. Kaplan,∫ Vladimir V. Tsukruk,§ Nancy Kelley-Loughnane‖‖ † UES Inc., 4401 Dayton-Xenia Road, Dayton, OH 45432, United States ‡ The Henry M. Jackson Foundation, 6720A Rockledge Drive, Bethesda MD 20817, United States § School of Materials Science and Engineering, Georgia Institute of Technology, 771 Ferst Drive NW, Atlanta, GA 30332, United States ∫ Department of Biomedical Engineering, Tufts University, 4 Colby Street, Medford, MA 02155, United States ‖‖ Air Force Research Laboratory, 711th Human Performance Wing, Airmen Systems Directorate, 2510 Fifth Street, Wright-Patterson AFB, Dayton, OH 45433, United States

Abstract Strategies for the encapsulation of cells for the design of cell-based sensors require efficient immobilization procedures while preserving biological activity of the reporter cells. Here, we introduce an immobilization technique that relies upon the symbiotic relationship between two bacterial strains: cellulose-producing Gluconacetobacter xylinus cells; and recombinant Escherichia coli cells harboring recombinase-based dual-color synthetic riboswitch (RS), as a model for cell-based sensor. Following sequential co-culturing of recombinant cells in the cellulose matrix, final immobilization of E. coli cells was completed after reconstituted silk fibroin (SF) protein was added to a “living membrane” generating the composite bacterial cellulose-silk fibroin (BC-SF) scaffold. By controlling incubation parameters for both types of cells, as well as the conformations in SF secondary structure, a variety of robust composite scaffolds were prepared ranging from opaque to transparent. The properties of the scaffolds were compared in terms of porosity, water capacity, distribution of recombinant cells within the scaffolds matrix, onset of cells activation, and ability to protect recombinant function of cells against UV irradiation. The closer-fitted microstructure of transparent BC-SF scaffolds resulted in leakage-free encapsulation of recombinant cells with preserved RS function due to a combination of several parameters that closely matched properties of a biofilm environment. Along with proper elasticity, fine porosity, capacity to retain the water, and ability of SF to absorb UV light, the composite hydrogel material provided necessary conditions to form confined cell colonies that modified cells metabolism and enhanced their resilience to the stresses induced by encapsulation. Key words: Bacterial cellulose, silk fibroin protein, composite scaffolds, recombinant E. coli cells, dual-color riboswitch, quorum sensing *Corresponding author: [email protected]

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INTRODUCTION Encapsulation of cells and specifically, microbial communities, has attained considerable interest in the field of biotechnology, as combinations of different fabrication processes and biomaterials continues to be explored.1-6 The variability in materials-processing design for the immobilization of living cells is based upon the application criteria and is primarily aimed to preserve cells viability, while allowing control of interfacial properties of living cells with the surrounding matrix.7,8 Several parameters must be taken into account for successful immobilization of active microbial cells, such as optimal choice of a biocompatible material host and use of appropriate immobilization method.7 If both of the parameters are satisfied, the proper entrapment of cells in biocompatible matrix will extend cells storage life, support their sensing and interrogation functions, provide access to signaling molecules, offer protection against environmental variables, and reduce the chance for environmental contamination by engineered microorganisms.9 The principle of cell encapsulation is based upon isolation of cells in semipermeable artificial membranes. This process can be accomplished via surface modification of individual cells, or bulk entrapment of clusters of cells. When modified with biocompatible polymers, immobilization of individual living cells, termed “artificial cells”, offers selective permeability, functional performance, biosafety, and chemical and mechanical protection.10 However, due to the nanothickness of polymer films, the effectiveness to sustain long-term storage of “artificial cells” may be compromised.11 Microencapsulation, on the other hand, provides a number of functional advantages over cell suspensions. Larger volumes of packing space can enhance protection against external forces during cultivation12 and provide defined microenvironments in order for cells to maintain their intrinsic properties.13 Using natural and synthetic polymers during microencapsulation, cells can be safely incorporated inside hydrogel biomaterials that provide many of the properties characteristic to external environments, such as viscoelasticity, mechanical and chemical stability, diffusion properties and nanofibril network architectures.14-16 Cellulose, a natural biopolymer, represents a versatile biomaterial that is widely used in biomedical applications due to its unique physical properties.17 Biocompatibility, excellent mechanical strength, high water absorption capacity and surface area, highly crystalline and refined fiber network structure render biocellulose useful in drug delivery and encapsulation systems for living tissues, cellular organisms, enzymes and other types of biocatalysts.18-22 Aside from traditional


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incorporation of cells in cellulosic scaffolds, the formation of “living membrane” systems that utilize symbiotic relationships between two types of bacterial organisms represents a novel biomimetic approach for living cell immobilization.23,24 For example, incorporation of Escherichia coli bacterial strains in cellulosic membranes produced by Gluconacetobacter xylinus was demonstrated to be an effective way to entrap recombinant cells during co-cultivation of both types of organisms.23 The compatibility of cellulose-producing bacteria with recombinant strains allows the development of “living membrane” systems that can be resistant to environmental degradation and persistent for short or long periods of time, making them durable and relatively easy to use for various detection applications. Recent advances in cell biology and biotechnology have provided an improved understanding of the complex dynamic interactions between cells and the host material, leading to new, useful functions.14 For example, novel living hybrid materials can help reporter cells implement functions as biosensors or bioreactors. A promising application of such hybrid systems is linked to the ability of bacteria to sense their environment with high sensitivity and selectivity.25 With progress in genetic manipulation and synthetic biology, living microbial cells can be engineered to detect specific analytes and produce a detectable signal, serving as diverse reporters in the field of medicine, environmental monitoring, food processing and safety.26-29 All of these biotechnological applications require long-term maintenance of metabolically active cells and fine control over their environment. Microenvironment defined by the host material has a major impact on cell behavior, as it can directly change cellular metabolism to resist harsh conditions, or indirectly relieve environmental stresses by maintaining proper cell homeostasis.30 When confined in limited space, individual bacterial cells communicate via quorum sensing (QS) signaling, a cell-to-cell communication system that for a long time was believed to be a prominent feature characteristic to high cell density populations.31,32 The information the bacteria perceive via different QS systems helps to assess nutrient availability, adjust a growth rate and coordinate collective behavior in order to produce the optimum adaptive capability for the maximum range of situations.31,33 This adaptation often involves changes in metabolism, motility, extracellular polysaccharide synthesis, biofilm formation, and similar processes important in virulence.34,35 Recently, it has been demonstrated that enough QS can occur in physically confined, isolated single cells and small populations to modify their metabolism and enhance cell resistance to



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external stresses.36-39 Hence, providing spatial confinement to individual, metabolically-active cells by restricting them in a proper hydrogel matrix with properties closely matching extracellular microenvironments is a promising biotechnological tool to develop robust and functional cellbased biosensors. In this study, we describe the formation of a composite “living membrane” system consisting of a bacterial cellulose (BC) matrix supplemented with silk fibroin (SF) protein and recombinant bacterial cells that harbor a theophylline synthetic riboswitch as a model biosensor. By adjusting the composition ratio of SF in BC host matrix, a variety of BC-based scaffolds were prepared that varied in optical transparency, mechanical stability, porosity, and protection against UV light. Particularly, the composite matrix provided a microenvironment that helped to avoid uncontrolled proliferation and facilitated close-fitting retention of cells in the host hydrogel matrix. Owing to their inherent similarity to the extracellular matrix and mild processing conditions, the composite matrix derived from natural polymers provided conditions for the microbial cells to maintain their recombinant functions as reporter cells. On the whole, composite BC-SF matrices with higher content of SF demonstrated direct physical protection of cells from harsh environmental conditions (UV light) and also indirectly resulted in the upregulation of QS signaling to enhance bacterial stress resistance.

RESULTS AND DISCUSSION Immobilization of one type of bacterial cell in a biocompatible matrix produced by another type of bacteria offers a simple and natural way to encapsulate cells of interest and provide physical protection. In this study, recombinant E. coli cells harboring a synthetic riboswitch were encapsulated in the native cellulose matrix produced by G. xylinus cells. The goal of this study was to develop an environmentally robust and long-term persistent “living membrane” consisting of BC matrix and recombinant bacterial strains entrapped within individual cellulosic sheaths. Since the recombinant E. coli cells were constructed to host a genetic cellular switch to detect the presence of relevant molecular targets, complete coverage of the recombinant cells and protection of their function were critical for developing a “living cell sensor”. Although, BC offers remarkable porosity, water absorbency and excellent biological affinity for sustaining the growth


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of secondary organisms, the permeability of the nanofibril network allows planktonic cells to escape from the BC matrix. To overcome this shortcoming, BC pellicles were saturated with a second natural polymer, reconstituted SF protein, forming composite bacterial cellulose–silk fibroin (BC-SF) scaffolds. This approach allowed to reduce the porosity of BC matrix and prevent the escape of the recombinant cells. Herein, we produced and compared several BC-based composite scaffolds that differed in BC and SF composition to examine the compatibility and properties of composite matrix to support the growth and function of the recombinant cells. Moreover, due to well-known natural protective and UV blocking properties of SF protein, composite BC-SF scaffolds were analyzed for their ability to shield and protect the recombinant bacteria from UV radiation. This will allow the composite substrates to be used as a protective immobilization matrix to enhance performance stability of cell-based biosensors in field-related applications.

Formation of pure BC and composite BC-SF scaffolds Figure 1 outlines the process for entrapping recombinant E. coli cells in either pure BC or composite BC-SF scaffolds, where several procedural steps required optimization. Incubation time necessary to produce the initial cellulose matrix, the amount of E. coli, the incubation period during co-culturing of both types of bacterial strains, the amount of SF solution, the viscosity and processing conditions required to initiate sol-gel transitions in silk fibroin structure, were all tested

Figure 1. Scheme outlining the formation of primary BC matrix and encapsulation of recombinant E. coli cells in pure BC or composite BC-SF matrix.



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to attain robust composite scaffolds with efficient entrapment of the recombinant cells. Table S1 refers to the incubation parameters that were optimized for producing pure and composite pellicles. Initially, the formation of stable BC pellicles had to be established, which required growing a single colony of G. xylinus cells for 3 days. Typically, during static cultivation, the cellulose matrix develops into a continuous polymer network of individual membranes accumulated at the airliquid interface.23 The pellicle thickness was directly proportional to the time of cultivation and was mainly limited by media carbon source and level of oxygenation.40,41 Followed the formation of the initial BC matrix (typically 3 days), the pellicles were inoculated with recombinant E. coli cells and co-cultured for another 3 days. This approach allowed a symbiotic relationship to be established between both types of cells and ensure homogeneous distribution of E. coli cells within developing cellulose network. Composite BC-SF scaffolds were produced by adding an aqueous solution of SF to already established colonies of E. coli cells (typically after 1 day of inoculation) to yield two types of composite BC-SF scaffolds: opaque and transparent. Opaque composite BCSF scaffolds were generated by adding 3-3.5% w/w aqueous SF solution in the ratio of 0.6:0.4 BCSF (w/w) and incubated for additional 2 days (30°C). Transparent composite BC-SF scaffolds were formed by adding 7-7.5% w/w aqueous SF solution in the ratio of 0.4:0.6 BC-SF (w/w) followed by incubation for 28 hours (39°C). Processing SF during continuous growth of recombinant cells in BC-SF matrix had a significant effect on the physical properties of the composite scaffolds. By controlling drying rate and water content in BC-SF matrix, composite scaffolds were produced with different mechanical and optical properties (Figures 2, S1). Figure 2a depicts the differences in absorbance between pure BC, and composite scaffolds with comparable amounts of E. coli cells loaded to pre-formed BC matrices. Regardless of the scaffold composition, there was a linear correlation between E. coli loading and optical properties of the scaffold matrix, demonstrating decreased transparency with increased amounts of E. coli cells (Figure 2b). The opacity of pure BC scaffolds was related to the continuous mass accumulation (both cells and cellulose) during longer incubation periods (3 days). The variability in physical properties of composite BC-SF scaffolds can be explained by the differences in conformational changes of SF secondary structure during sol-gel processing. Degradable opaque BC-SF composite scaffolds were formed as the result of formation of metastable supramolecular fibroin structures comprised


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of an amorphous bulk structure (random coils, α helixes, β turns and bends) with a low content of β sheets.42 On the other hand, water-stable transparent BC-SF scaffolds were formed via conditions closely matching water annealing.43,44 These scaffolds were reported to have higher thermodynamically stable silk II content (antiparallel β sheets) than silk I content (hydrated type II β-turns), and were translucent, held more water and were temperature stable.43,44 Microstructurally, pure BC and composite BC-SF scaffolds also differed significantly. Figure 3 illustrates the morphology of the nanofibral network arrangement in lyophilized BC and BC-SF scaffolds and provides the distribution of pore sizes for each type of the scaffold. Assessing the pore sizes in all types of the scaffold matrices allowed us to determine the optimal design to provide complete entrapment of the cells, as well as to predict permeability of analyte molecules. Although direct correlation between hydrated and lyophilized materials structure cannot be drawn,



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the SEM imaging provided information on the structural arrangement of the fibers in each of the matrices and allowed an estimate of their porosity. Pure BC scaffolds formed a continuous 3-D network of intertwined cellulosic nanofibrils with void spaces exceeding the length of rod-shaped E. coli bacteria (typically 2 µm in length) (Figure 3ab). The pore sizes of a mesh-like network structure were 8.4 ± 2.5 µm and this is likely to be an underestimation of their size in fully hydrated state. A high permeability of pure BC scaffolds caused leakage of motile recombinant cells from the matrix, as will be discussed later. On the other a)

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hand, composite BC-SF scaffolds formed heterogeneous network structures where cellulose nanofibrils were covered by SF solution (Figure 3c-f). During formation of BC-SF nanocomposite scaffolds with varying amounts of SF, increased concentration of SF led to decreased porosity. The mesh size of the network was reduced below 1 µm for BC-SF scaffolds with highest SF concentration (7% w/w) (as estimated from dehydrated samples) (Figure 3e-f). This input minimized the loss of recombinant cells to the surrounding media and ensured complete retention of E. coli cells, particularly in the transparent BC-SF composite scaffolds. Water absorbency, water retention and mechanical properties Measurements of the swelling ratio and water absorption capacity were performed after incubation of the scaffolds in DI water for 24 hours. Pure BC scaffolds were highly saturated with water (94% of moisture content) and demonstrated up to a 14-fold increase in swelling ratio from its dry weight (Figure S2a-b). Over 90% of the total water content was absorbed within 30 min, emphasizing the highly porous nature of the BC scaffolds. The reduction in water uptake capacity for the composite BC-SF scaffolds was correlated to increased SF content. In composite BC-SF scaffolds, a)

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Figure 4. AFM topography images of pure BC (a) and transparent BC-SF (b) scaffolds taken under ambient condition. Comparison of apparent Young’s modulus (in swollen state) for pure BC and composite opaque and transparent BC-CF scaffolds (c).



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the swelling ratio decreased to about half of that for the BC scaffolds, however the water content declined only slightly. Opaque BC-SF scaffolds had a 6.4-fold increase in swelling ratio with a 91% water content. Transparent BC-SF scaffolds showed a 4.6-fold swelling ratio and 86% water content. AFM images showed morphology of BC and BC-SF network (at ambient conditions) with integrated cells (Figure 4a-b). Stiffness of pure BC matrix (in wet state) was derived from the force distance curves obtained with AFM indentation using colloidal probes (see Experimental).45 The results were analyzed with a Hertzian contact model to give an effective “apparent” modulus of 16.1 ± 9.2 kPa, which shows low compression stress resistance of pure low-density network of cellulose nanofiber morphologies (Figure 4c). The composite BC-SF scaffolds had significantly higher compression resistivity, with an increase in apparent Young’s modulus by three orders of magnitude in comparison to pure BC matrix. In particular, indentation results indicated a modulus of 9.6 ± 1.9 MPa for the opaque and a modulus of 13.4 ± 3.6 MPa for the transparent scaffolds with both values being comparable to common rubbery materials (Figure 4c).45 The apparent decrease in water absorbency and gain in elastic properties were due to the increased SF content. As it can be seen in the AFM topography results obtained from the scaffolds (Figure 4a-b), the addition of silk fibroin causes much of the loosely packed cellulose matrix to be filled with silk, thereby reducing the amount of space accessible to incoming water. It has previously been demonstrated that the modulus of a scaffold has a direct impact of the speed of cellular migration through the scaffold material.46 In addition, there have been several reports demonstrating the effect of scaffold stiffness on the stem cell differentiation fate for mesenchymal stem cells.47,48 Composite materials have been shown previously to be an effective method of enhancing scaffold modulus for altering cellular gene expression.49 The probe size and forces used in the indentation measurements here closely match those of E. coli cells that have been previously studied, thus the stiffness measurements obtained here are likely good approximations of the stiffness observed at the cellular level.50 Activation of dual-color riboswitch in E. coli cells Recombinant E. coli cells utilized in this study were transformed with a bidirectional reporter system, consisting of recombinase FimE controlled by theophylline synthetic riboswitch and an


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invertible DNA segment containing a constitutively active promoter placed between two fluorescent protein genes, GFPa1 and mKate2.51 This system allowed us not only to localize and assess the fitness of encapsulated E. coli cells, but also to monitor the occurrence and propagation of fluorescence signal across the living membrane. In the absence of a target analyte (theophylline), cells expressed GFPa1 protein and hence appeared green and viable. However, when the analyte was present, the activation of FimE recombinase facilitated the inversion of the promoter leading to constitutive expression of mKate2 protein, causing the cells to appear red. Co-cultivation of cellulose producing bacteria with secondary catalytic organisms led to the development of a compatible “living membrane”. Continuous accumulation of E. coli in G. xylinus supportive media not only demonstrated the compatibility of the present technique to entrap secondary organisms, but also allowed to achieve flexibility in maintaining baseline populations. Since it was not possible to directly estimate the density of recombinant cells in the final pellicles, the dual-color reporter system allowed us to qualitatively assess the amount of viable E. coli cells based on the level of GFPa1 intensity. The number of E. coli cells added to the growing cellulose pellicles had a direct correlation on the kinetics of cell growth and hence the level of expression for fluorescent proteins. GFPa1 fluorescence intensity showed dose-dependent increase by ~6-fold when the seeding number of E. coli cells was increased from 2.5∙107 to 1.5∙108 (Figure 5a). Hence, with the higher number of entrapped cells, the higher level of inducible signal was achieved. On the other hand, with increased loading of E. coli cells (typically above 109 cells), the crossover in color predominance between non-activated and activated cells was delayed, limiting the sensitivity of the recombinant cells to detect the target analyte. Thus, the optimal loading number of cells was set between 5∙107 and 1∙109 cells to balance the two signals. As a measure of the effective diffusion of small solutes, the scaffolds were tested for kinetics of cells expression, since inducer molecules have to diffuse and induce the expression of fluorescent proteins. The progression of inducible mKate2 signal and decrease in production of GFPa1 protein were monitored from the whole pellicles. The fluorescence signal was averaged from several locations across each pellicle to give an objective comparison in cell fitness related to the material. The ability of recombinant cells to detect the target analyte and produce fluorescent signal while being encapsulated in various cellulose-based scaffolds was also analyzed using two cell loading



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numbers: 3.3∙108 and 1.5∙108 (Figure 5b-d). The fluorescence levels were not only sensitive to the seeding number of recombinant bacteria, but also to the type of scaffold cells were entrapped in. Particularly, activation of E. coli cells in pure BC membranes demonstrated rapid expression of mKate2 protein, with the majority of cells producing red fluorescent protein after 20 or 24 hours of incubation for the 1.5∙108 or 3.3∙108 cell loading numbers, respectively (Figure 5b). E. coli cells encapsulated in opaque BC-SF scaffolds showed a gain in fluorescence intensity after 30-35 hours of incubation, while cells encapsulated in transparent BC-SF scaffolds showed predominance in red fluorescence only after 20-22 hours of incubation (Figure 5c-d). The apparent difference in a)

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the delay and reduction level of mKate2 protein expression in E. coli cells encapsulated in opaque BC-SF scaffolds was not only due to decreased diffusion, but also related to the opacity of composite material, which is a major constrain of highly diffracting material network in signal detection. The high porosity of pure BC scaffolds, along with large amount of water entrained within cellulosic membranes provided fast diffusion for analyte molecules to reach the recombinant cells and elicit the response. The mechanism of diffusion of small hydrophilic molecules in pure BC membranes has been mainly attributed to the pore sizes with some hindering effect due to the fiber obstruction.18,52 We also predict that the partitioning of small solutes (analyte molecules) with recombinant cells (analyte responsive) will affect the overall diffusion, particularly for the scaffolds containing high densities of cells. Although, for highly swollen substrates (pure BC scaffolds), the pore mechanism will likely prevail in the transport of solute molecules with low molecular weight. For less permeable substrates, such as in the case of composite BC-SF scaffolds, the hindering effect of smaller effective void dimensions in the matrix mayl account for reduced diffusion caused by hydrodynamic and entropic restrictions. From a materials point of view, pure BC scaffolds promoting fast activation of cells might seem optimal for the storage of recombinant cells. However, during incubation in the reaction buffer, E. coli cells were able to easily escape from highly porous cellulosic membranes under sheer forces. As illustrated in Figure 3, the microstructure of cellulose membranes consists of loosely intertwined nanofibrils with the mesh pore sizes much greater than the size of recombinant cells. This highly porous scaffold supported continuous efflux of E. coli cells into the media where they then proliferated, causing an increase in fluorescence signal and optical density. Figure 6 provides kinetics for both types of fluorescent proteins (GFPa1 and mKate2) and of cell growth measured from planktonic E. coli cells accumulated in the media. The results show continuous increase in mKate2 fluorescence and growth patterns over the time as the cell loading number increases (Figure 6a-b). On the other hand, when the recombinant cells were encapsulated in the composite BC-SF scaffolds, the cell escape was significantly reduced only from the transparent BC-SF matrix (Figure 6e-f). Opaque BC-SF scaffolds demonstrated a modest decrease in efflux of planktonic cells due to the specific microstructure of the composite matrix (Figure 6c-d and Figure 3c-d).



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The processing of low viscosity SF during the formation of opaque BC-SF pellicles reduced pore dimensions insufficiently to impede the leakage of the cells (Figure 3c-d). With a higher content of SF in the composite BC-SF matrix, a denser network structure was formed, reducing the effective voids (Figure 3e-f) and hindering the escape of E. coli cells from BC-SF scaffolds (Figure 6e-f). Hence, only the transparent composite BC-SF scaffolds (with higher content of SF)


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provided the most effective entrapment matrix that both supported catalytic activation and hindered diffusion of recombinant cells. Distribution of activated E. coli cells in cellulose-based scaffolds Figure 7 illustrates the differences in spatial distribution of recombinant cells within pure and composite scaffolds after continuous cultivation for 48 hours. Spatial fluctuations of fluorescence intensities were indicative of differences in density of E. coli cells. As shown in localized confocal images, activated E. coli cells were radially distributed across pure BC pellicles, with the central a)

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Figure 7. Distribution of E. coli cells encapsulated in pure BC (a) and transparent BC-SF (b) composite scaffolds after incubation in the reaction media for 48 hours (2.5 mM theophylline, DMSO, 37°C, 110 rpm). Optical fluorescent image of the whole pellicle was combined with three confocal images from the selected areas. Red fluorescence was attributed to E. coli cells expressing inducible mKate2 protein (activated cells), green fluorescence was representing E. coli cells constitutively expressing GFPa1 protein (non-activated cells) and blue fluorescence was associated with intrinsic fluorescence of SF.



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region having the lowest cell density and the periphery of the pellicles the highest (Figure 7a). Such behavior can be explained by the continuous influx of actively proliferating motile bacteria back into the BC matrix. Due to the highly porous microstructure of the pure BC network, inward propagation of cells led to a gradual saturation of the pellicle circumference with activated cells. In contrast to pure BC scaffolds, the final localization of recombinant cells in the composite BCSF scaffolds occurred during sol-gel processing of SF, where recombinant cells were restricted in the movement due to reduced porosity of the matrix (Figure 7b). The recombinant cells had an even distribution across the composite matrix with some exceptions in the form of small localized cell colonies. Since the porosity of the composite matrix was less than the effective radii of the recombinant cells, the formation of local populations might have been occurred during the processing of SF. With time permitted to form several generation of bacterial cells (processing of SF occurred within 10 hours), small populations of E. coli cells were formed, which later were confined to the spaces limited by excess of SF. The ability of cells to undergo division during SF processing illustrates the compatibility of the current technique for cell encapsulation process. Hence, composite BC-SF matrices with a higher SF content (~7% w/w) provided preferable conditions for colonization and aggregation of bacteria, and for the isolation and activation of recombinant cells. Protection of BC-SF scaffolds against UV light Whole cell-based sensors utilizing recombinant microorganisms are more sensitive to harsh environmental conditions than conventional sensor devices.53 In particular, prolonged exposure to ambient sunlight intensity can be lethal to microorganisms lacking physical protection mechanisms against UV radiation.54,55 Thus, developing biomimetic approaches that can enhance protection against UV-related damage is essential for many biotechnological applications requiring stable performance of engineered microbes.56 It has been know that silk can absorb UV light due to its tyrosine-rich amino acid composition.57 However, whether a specific primary structure of polypeptide arrangement has more prominent effect on UV absorption has never been investigated.58 Here, we compared the effectiveness of reconstituted silk fibroin in its various forms (predominant silk II structure in transparent BC-SF scaffolds and amorphous structure in opaque BC-SF scaffolds) to effectively absorb UV light and


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protect recombinant cells from inactivation. BC-based pellicles (both pure and composite) containing comparable cell loading numbers were irradiated with handheld UV lamp for 2 hours (λ=254 nm, dose of irradiation was 380 J∙cm-2). Time-course studies revealed different levels of bacteria susceptibility to UV radiation under conditions exceeding the recommended doses for inactivation of E. coli bacterial populations (~36 mJ∙cm-2).59 Particularly, UV-treated suspensions of bacterial cells (negative control) did not proliferate and lost their ability to react to an inducer analyte over the course of study (Figure 8a). Similarly, E. coli cells encapsulated in pure BC scaffolds lost their ability to respond to the presence of analyte and did not show RS activation (Figure 8b). In contrast, E. coli cells encapsulated in composite BC-SF scaffolds demonstrated response recovery, however, not to the same extent as non-UV treated cells immobilized in the same type of matrix. Even though the activation of cells was initiated at the same time point, the response rate was significantly slower. The gain in fluorescence intensity from the recombinant cells encapsulated in opaque BC-SF composite scaffolds was reduced by 80% after 52 hours of post-induction compared to non-irradiated scaffolds (Figure 8c). Maximum fluorescence for UVirradiated E. coli cells encapsulated in transparent BC-SF scaffolds reached about half of that for the non-UV treated scaffolds (Figure 8d). Two factors were attributed to the bacterial survival in BC-SF scaffolds: the ability of SF to absorb light in UV-C region and the confinement of E. coli cells in the composite matrix. Absorption of SF protein in 200-280 nm range is accredited to aromatic amino acids residues, tyrosine (Tyr, 5.3 mol.%), tryptophan (Trp, 0.5 mol.%) and phenylalanine (Phe, 0.6 mol.%) that have distinguishable absorption peaks at 228 and 278 nm (Figure S3).60 Increased β-sheet structure during the formation of solidified silk fibroin films can enhance optical responses further.61 Such an effect was attributed to stronger intra- and inter-molecular hydrogen bond interactions including the aromatic amino acids, specifically π→π* charge transfer transitions of Tyr and Trp aromatic rings.62 We observed significant enhancement in absorption peaks for water vapor annealed SF cast films compared to the corresponding SF solutions that also linearly correlated to SF concentration (Figure S3). Additionally, during the formation of composite BC-SF scaffolds with



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Figure 8. Comparison of RS activation in E. coli cells after UV irradiation (254 nm, 3.8 MJ∙m-2). Nonencapsulated E. coli cells (control) (a) were cross-referenced with cells encapsulated in pure BC (b), opaque BC-SF (c) and transparent BC-SF (d) scaffolds. Kinetics of fluorescence accumulation in pellicles (for both GFPa1 (green lines) and mKate2 (red lines) proteins) were compared to optical images captured at specified time points. Cells were induced with 2.5 mM theophylline (100 mM, DMSO) and incubated in LB media (37 °C, 110 rpm). (Data represent average ± SD (n=8) from two independent experiments).

higher SF content (7% w/w), water-annealing treatment might enhance secondary structure towards thermodynamically stable β-sheet conformations, increasing silk II content.42,44,61 Hence,


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by inducing β-sheet structure during the formation of composite scaffolds, UV–light absorption or blocking can be enhanced, enabling the recombinant cells encapsulated in high content SF composite scaffolds to have improved protection against UV exposure. In addition to UV absorption, the protective potential of composite BC-SF scaffolds was indirectly attributed to promoting clustering of E. coli cells. Localized colonies of recombinant cells were formed during the preparation of the composite scaffolds (Figure 7b). Establishment of cell colonies provided additional protection to bacterial fitness, enhancing their metabolic capacity and improving survival during stress, which will be discussed in the next section. Quorum sensing capacity of recombinant E. coli cells Understanding the physiological state at which the entrapped recombinant cells initiated the expression of plasmid-encoded genes is important for the design and characterization of cell-based sensors. Usually, the metabolic state of individual bacterium is communicated via quorum sensing (QS) signaling and can be coordinated among cell populations.63 Here, we evaluated the metabolic potential of encapsulated E. coli cells to activate RS function and initiate expression of recombinant proteins, particularly when they were predisposed to UV irradiation. The production and release of an extracellular factor, termed autoinducer-2 (AI-2) was measured to assess the function of cells.31 Measuring AI-2 signaling was based on the auto-induction of bioluminescent response from Vibrio harveyi, a luminescent marine bacterium that senses and reports extracellular AI-2 levels from another bacterial species.64,65 The results in Figure 9 and Figure S4 depict kinetics profiles of AI-2 released from E. coli cells during activation of RS. Maximum AI-2 activity for non-UV exposed E. coli cells (positive control) was recorded at an early exponential cell growth phase (~5 hr) and then steadily declined over the course of study (Figure 9a). Simultaneously, the maxima for accumulation of AI-2 levels was correlated to the initiation of heterologous mKate2 gene expression (Figure 8a). UV-exposed E. coli cells (negative control) demonstrated significant inhibition in AI-2 activity that corresponded to repressed cell growth and loss of cellular activity after UV irradiation. In contrast, cells encapsulated in BC-based scaffolds demonstrated attenuated AI-2 signaling activity that correlated to the mechanical properties of the scaffolds (specifically, microporosity and SF content). Only a 15% decrease in maximum AI-2 activity (compared to positive control)



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was observed for BC-encapsulated E. coli cells that was linked to active proliferation of cells in highly porous cellulose scaffolds (Figures 7a and 9b). On the other hand, UV-exposed BCencapsulated cells demonstrated significant inhibition in AI-2 activity (90%) correlating to the inhibition of growth kinetic profiles (Figure 9b). Interestingly, attenuation of AI-2 activity profiles for cells encapsulated in composite BC-SF scaffolds demonstrated a 35-40% decrease in comparison to the positive control (Figure 9c-d). QS activity, in contrast, was not affected by the exposure to UV light when E. coli was encapsulated in transparent BC-SF matrices, and an additional 25% reduction in AI-2 levels was observed when cells were immobilized in opaque BC-


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SF matrices (Figures 9c-d). In all events, attaining maximum QS signaling preceded the initiation of RS activation and correlated as an indicator of the metabolic state of the recombinant cells. The synthesis and uptake of AI-2 is intrinsically linked to the metabolic potential of E. coli cells, particularly with regard to the regulation of gene expression under stress-related conditions.66 The assessment of QS signaling among recombinant cells encapsulated in various types of cellulosebased scaffolds allowed us to evaluate whether a “cellular burden” was associated with the confined environment and how physical isolation affected the ability of the cells to mitigate extreme conditions. The variability in kinetics of QS signaling profiles suggested that entrapment of recombinant cells in the BC-based matrices affected expression of heterologous genes to various degrees, defined by the properties of the scaffold. Thus, the pliable and highly porous structure of pure BC scaffolds provided favorable conditions for the secondary organisms to survive, proliferate and maintain their physiological properties, analogous to the positive control cells. This was supported by continuous efflux of recombinant cells from the pellicles, increase in cell density across the scaffolds and a decrease in AI-2 signaling. By normalizing the bulk data for AI-2 activity to a single cell unit, the kinetic profiles looked similar for both non-encapsulated and BCencapsulated cells (Figure S3a-b). In other words, cells had a low QS capacity when they were not physically and spatially restricted. Unlike the highly porous BC matrices, the more rigid and less porous microstructure of the composite BC-SF scaffolds led to hindered cell proliferation accompanied by the formation of small isolated populations spatially-localized. Such confinement resulted in upregulation of QS activity (if calculated for a unit cell) (Figure S4c-d). Specifically, it implied that isolated cell microencapsulation significantly improved QS capacity of the recombinant cells and benefited their metabolism. When comparing the metabolic potential of non-irradiated and UV-irradiated cells encapsulated in the tighter BC-SF matrices, the cells “reported” similar levels of QS signaling, suggesting that no additional burden was associated with the UV irradiation (Figure S4d). On the other hand, cells encapsulated in pure BC scaffolds suffered significantly after exposure to UV irradiation showing significant decrease in QS capacity (Figure S4b). Hence, the confined environment facilitated the upregulation of the metabolic potential and could enhance resistance to harsh environmental conditions. Together with the beneficial properties of SF to absorb UV light, a closer-fitting microstructure of composite BC-SF scaffolds improved



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encapsulation and storage of our model recombinant cells, as a prototype substrate for cell-based sensors. CONCLUSIONS Encapsulation of recombinant E. coli cells harboring dual color RS was performed in biocompatible composite scaffolds consisting of a bacterial cellulose matrix supplemented with reconstituted silk fibroin material. The ease and simplicity of the present technique relied upon the symbiotic relationship between both bacterial organisms, where one provided a polymeric matrix for immobilization and storage of the secondary organism. By varying the composition ratio between cellulose and the additive SF, as well as processing conditions and viscosity for SF during the formation of composite scaffolds, matrix properties can be controlled. The optimal encapsulation matrix was accomplished when higher concentration and content ratio of SF was used. It led to the formation of robust and transparent composite scaffolds with matrix properties closely matching extracellular environment. The closer-fitted, pliable and biomimetic composite scaffolds not only preserved recombinant function of cells, but also improved their metabolic potential. Particularly, confining cells in the form of spatially-localized populations led to the upregulation of metabolic pathways and improved QS capacity. This was particularly beneficial for bacteria to coordinate collective gene expression and adapt population-driven behavior, which usually involves changes in community properties and improved bacterial resistance to local microenvironments. When exposed to UV-light, cells were able to recover, respond and activate analyte-induced heterologous genes. Both factors, the spatially-confined environment and strong UV-absorbing properties of β-sheet structures in SF-rich scaffolds allowed E. coli cells to better maintain their recombinant properties and resist additional stresses associated with exposure to UV light. Adopting the present immobilization technique on a macroscale level can be beneficial for biotechnological applications that require optical transparency of the material host to be surveyed from a long distance.

ACKNOWLEDGEMENTS This research was funded by the Air Force Office of Scientific Research (AFOSR) Grant 16RH3003J through Natural Materials and Systems Program (Dr. Hugh DeLong Office); Research


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Associateship Grant FA9550-12-D-0001 from National Research Council; AFOSR Grant FA9550-14-1-0269 and the US National Science Foundation CBET-1401720 Grant. Partial funding was provided through the Applied Research for the Advancement of Science and Technology Priorities (ARAP) Program in Synthetic Biology for Military Environments Grant #0602251D8Z. We also appreciate Dr. Hugh DeLong for valuable comments and discussions.

Supporting Information Supplemental experimental procedures; images of BC-based pellicles; swelling ratio and moisture content; UV-Vis spectra of silk solutions and dry films; normalized AI-2 activity profiles for E. coli cells after irradiation with UV light.



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EXPERIMENTAL SECTION Materials. Luria-Bertani (LB) (Difco, Lenox) broth powder, Bacto trypton, yeast extract, Bacto b) agar were purchased from BD (Becton, Dickenson and Co.) (Sparks, MD). Sodium chloride, potassium chloride, sodium phosphate dibasic anhydrous, potassium phosphate monobasic, theophylline, ampicillin, chloramphenicol, anhydrous citric acid, dimethyl sulfoxide (DMSO) have been purchased from Sigma-Aldrich Co LLC (Milwaukee, WI). Corning Costar cell culture plates (12-well and 6-well, transparent Corning grade) have been purchased from Sigma-Aldrich Co LLC (Milwaukee, WI). Bacterial Cell Strains. Gluconacetobacter xylinus (G. xylinus) (Brown) 10245TM strain cells were purchased from ATCC (Manassas, VA). G. xylinus cells were rehydrated and cultured in 5 mL of Hestrin-Scharamm (HS) medium containing 5 g·L-1 Bacto trypton, 5 g·L-1 yeast extract, 2.7 g·L-1 anhydrous disodium phosphate, 1.5 g·L-1 citric acid and 20 g·L-1 glucose. Cells were cultured using 15 mL culture tubes (28ºC, 220 rpm). E. coli cells were transformed with a plasmid containing a bidirectional reporter switch, pHWG640:mKate2-fimS-GFPa1, and a plasmid expressing FimE recombinase under control of theophylline synthetic riboswitch (RS), pSAL:RSFimE.51 Cells were cultured in LB medium supplemented with ampicillin (100 µg/mL) and chloramphenicol (25 µg/mL) at 37°C, 220 rpm overnight before inoculation. Vibrio harveyi (V. harveyi) strain BB-170 was obtained from ATTC (Manassas, VA). Cells were n cultured in autoinducer bioassay (AB) medium at 30°C (170 rpm) using 15 mL culture tubes for 24 hours before assessing levels of autoinduction AI-2 from E. coli cells. Reconstituted Silk Fibroin. Solution of reconstituted silk fibroin (SF) was obtained from silk cocoons produced by Bombyx mori silk worms according to previously reported protocols.67,68 Briefly, cocoons were stripped on individual membranes were degummed with 0.02 M Na2CO3 solution at 100°C for 30 min and washed thoroughly with deionized water. Extracted silk fibroin fibers were dissolved in 9.3 M LiBr solution at 60ºC for 2 h, yielding a 20% (w/w) solution. The solution was dialyzed against deionized water using Slide-a-Lyzer dialysis cassette (molecular weight cutoff (MWCO) 3,500 Da, Pierce) at room temperature overnight. An aqueous SF solution free of impurities was obtained after centrifugation (3X, 20,000 rpm, 20 min, 4ºC). The final concentration of SF was ~7% (w/w) as determined by weighing of the dried solids. Lower concentration of SF (3%, w/w) was obtained by careful diluting 7% solution with deionized water. All stocks of SF solutions have been stored at 4ºC before use. Encapsulation of E. coli in Pure BC and Composite BC-SF Scaffolds. Pure bacterial cellulose (BC) scaffolds were formed by inoculating 5 mL fresh HS media with 50 µL six-day culture aliquots. Cultivation proceeded in a static mode using 12-well cell culture plates at 28ºC for various time intervals. This allowed to produce initial BC matrix consisting of individual BC membranes having diameter of 2.5 cm. To form BC scaffolds with encapsulated E. coli cells, aliquots (5 µL, 15 µL, 30 µ, 50 µL, 100 µL and 200 µL) of overnight culture of E. coli cells (37ºC, 220 rpm) have been added to the wells containing preformed BC matrix. Both cell cultures have been allowed to


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incubate at 37ºC for 3 days forming pure BC scaffolds that contain both types of bacteria, cellulose producing and recombinant E. coli cells. Composite bacterial cellulose-silk fibroin (BC-SF) scaffolds were produced by adding reconstituted aqueous silk fibroin solution to BC matrix containing E. coli cells after 1 day of inoculation. Opaque BC-SF scaffolds were formed using 0.5 mL of 3% w/w SF solution during slow incubation periods (48 h, 30ºC). Transparent BC-SF scaffolds were formed by using 0.3 mL of 7% w/w SF solution and incubating at 37ºC for 10 h followed by incubation in PBS solution for 18 h (1X, 0.5 mL). Prior to adding SF solution, excess media containing planktonic bacteria was removed. Alternatively, excess media was removed from the control plates - BC scaffolds with E. coli cells and substituted with PBS solution (1X, 0.5 mL or 0.3 mL), followed by incubation at either 30ºC for 48 h or at 37°C for 28 h to match conditions for corresponding BC-SF scaffolds. Samples were termed according to their opacity: opaque and transparent BC-SF scaffolds. Water-uptake Capacity. The swelling ratio was calculated by placing separately freeze-dried pure BC and composite BC-SF samples of 2.5 cm in diameter in distilled water for specified time intervals. Samples were weighed at 1 and 5 min, followed by measurements every 5 min up to 30 min, and then after 60, 180 and 1200 min. After removal from the distilled water, excess superficial water was removed by gentle tapping with filter paper. The content of the distilled water in the swollen scaffolds was calculated by the following equation: water uptake (%): [(Ws – Wd)/Ws] × 100%, where Ws and Wd are the weights of the swollen and the dry scaffolds, respectively. All experiments were performed in triplicates and standard deviations were calculated. Mechanical Testing of BC-based Scaffolds. All AFM images were collected using standard AFM tips with radius ~8 nm (MikroMasch). Imaging was performed in tapping mode using either a standard tip holder for ambient conditions or a fluid cell for scanning in liquid. Mechanical testing was performed through nanoindentation experiments via an atomic force microscope (Dimension Icon, Bruker).45 All samples were mounted on double sided tape and scanned under ambient conditions before testing. Mechanical testing was performed in deionized water using a fluid cell, and 10 force distance curves were obtained for each sample. These force distance curves were analyzed with a Hertzian contact model (using NanoScope Analysis) to obtain average (n =10) values for the apparent Young’s modulus of the scaffolds. The mechanical data for all samples was collected with colloidal borosilicate AFM probes (Novascan) with a 1.25 µm radius and 14.2 N/m spring constant. Much larger contact area of colloidal probes allows for tip interaction with whole network, not individual fibrils, thus reflecting stiffness of the entire materials structure. The pure bacterial cellulose scaffold required the use of a probe with 5 µm radius and 0.0248 N/m spring constant due to the their very low stiffness. Considering highly porous structure of BC network, the contact areas cannot be estimated correctly within Hertzian model, and thus, the absolute values of the elastic modulus reported here represent only apparent values related to compression resistance/stiffness of the composite materials. The deflection sensitivity of the cantilever was determined before testing through deflecting the probe on a



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sapphire standard under fluid conditions. The spring constants of the cantilevers were determined via the thermal noise method after evaluation of the deflection sensitivity. Activation of E. coli Cells. Cells were induced with 2.5 mM of theophylline (100 mM stock solution, DMSO) while incubation was performed in LB media (37ºC, 110 rpm) using 6-well cell culture plates (Corning). Prior to activation, all scaffolds were briefly washed in PBS solution to remove any un-immobilized cells. Fluorescence Intensity Measurements. The time course of E. coli cell activation was performed on a SpectraMaxM5 multimode microplate reader (Molecular Devices, LLC, Sunnyvale, CA) using 12-well cell culture plates. Scaffolds were removed from 6-well cell culture plates and briefly washed in PBS buffer prior to measurements. Excitation at λex=470 nm and λex=588 nm was used for monitoring GFPa1 and mKate2 fluorescence, respectively. Emission peaks at λem=510 nm and λem=620 nm were used to characterize fluorescence for GFPa1 and mKate2 proteins expressed by cells, respectively. Absorbance of the media was acquired at λ=600 nm to monitor the release of escaped cells from BC and BC-SF scaffolds, once the pellicles were removed from the plates. UV-Vis Extinction Spectra. UV-Vis spectra were obtained on SpectraMaxM5 microplate reader using 12-well cell culture plates (transparent Corning). Absorption spectra in transmittance mode were collected from wet samples of pure BC and composite BC/SF scaffolds (n=5). Data was averaged and normalized to the empty plates. Experiments were performed using degree of freedom n-1=4, standard deviations were calculated. Scanning Electron Microscopy (SEM). Imaging has been performed on Jeol-6610LV system at 2.5 kV accelerating voltage. Prior to imaging, samples were lyophilized for 24 hours using LABCONCO FreeZone freeze dry system. Lyophilized samples were carefully cross-sectioned with a razor blade and mounted to a conductive adhesive tape on copper stubs. Thin gold films (~3-5 nm) were sputtered on the samples prior to imaging. Porosity Study. Porosity was analyzed by two complimentary methods - analyzing pore sizes from SEM images using ImageJ software and performing molecular weight cut-off (MWCO) exclusion limit study using FITC-labeled dextrans of variable Mw. The evaluation of pore sizes was performed by examining several SEM surface and cross-section images of lyophilized pure BC and composite BC-SF samples and performing average calculations. The calculations were based on averaging the pore sizes from samples of 600 x 800 µm2 and by calculating coefficient of variation with α=0.95 confidence level. Fluorescent Imaging. Optical images of fluorescent scaffolds were obtained with FujiFilm FinePix camera using UV filter during illumination at λex=400-500 nm on dark transilluminator (Clare Chemical Research, Dolores, CO). Confocal imaging was performed on Olympus FV1000 system using 405 nm, 488 nm and 543 nm lasers.


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UV Exposure. Control E. coli cells (100 µL in PBS), BC and BC-SF scaffolds containing E. coli cells were exposed to short range of UV light (λ=254 nm) by irradiating with UVGL-58 handheld UV lamp (UVP, LLC) from a distance of 3 cm for 2 hours. The pellicles were flipped over after 1 hour of irradiation to achieve full exposure to UV light. The dosage of UV irradiation was calculated to be 380 J∙cm-2. Quorum Sensing and AI-2 Detection. The detection of autoinducer-2 (AI-2) was performed with V. harveyi cells. Over the course of study, 100 µL culture supernatant from the reaction media where various BC-based pellicles were n incubated was collected and centrifuged at 8,000 rpm for 10 min and filtered with sterile 0.22 µm filter system during centrifugation at 13,000 rpm. Cellfree supernatants were stored at -20°C for no longer than 2 days. In the detection assays, overnight cultures of V. harveyi cells were diluted at 1:5000 into fresh AB medium. In 96-well plates (Corning brand), 180 µL of diluted culture of V. harveyi cells was combined with 20 µL of cellfree culture supernatant from experimental samples and incubated at 30oC for 4 hours. For negative controls, 20 µL of sterile AB media was used. For positive controls, 20 µL of cell-free supernatant from V. harveyi culture was used. The bioluminescence intensity was measured by SpectraMaxM5 microplate reader in the luminescence mode. Each sample was used in triplicates, and each data point represents three independent samples. The AI-2 production was quantified as relative AI-2 concentration and was calculated as follows: [Nt0/Np] × 100%, where Nt0 = Nt – N0 represents true bioluminescence of the experimental sample, Nt represents crude bioluminescence of the sample, N0 is the bioluminescence of negative control and Np is bioluminescence of positive control.



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For Table of Contents Use Only

Immobilization of Recombinant E. coli cells in a Bacterial Cellulose-Silk Composite Matrix to Preserve Biological Function Irina Drachuk, Svetlana Harbaugh, Ren Geryak, David L. Kaplan, Vladimir V. Tsukruk, Nancy Kelley-Loughnane Synopsis: Macroscale immobilization of dual color recombinant E. coli cells in a composite matrix consisting of bacterial cellulose and reconstituted silk fibroin protein.


Immobilization of Recombinant E. coli Cells in a Bacterial Cellulose

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