An in vitro tissue model for screening sustained release of phosphate-based therapeutics in attenuating pathogen-induced proteolytic matrix degradation

Abstract

Tissue response to intestinal injury or disease releases pro-inflammatory host stress signals triggering microbial shift to pathogenic phenotypes. One such phenotype is increased protease production resulting in collagen degradation and activation of host matrix metalloproteinases contributing to tissue breakdown. We have shown that surgical injury depletes local intestinal phosphate concentration triggering bacterial virulence and that polyphosphate replenishment attenuates virulence and collagenolytic activity. Mechanistic understanding of how bacterial and host protease expression contributes to tissue breakdown is difficult to achieve in vivo necessitating the development of novel 3D in vitro tissue models. Common techniques for screening in vitro protease activity including gelatin zymography or fluorogenic protease-sensitive substrate kits do not readily translate to 3D matrix degradation. Here, we report the application of an in vitro assay in which collagenolytic pathogens are cultured in the presence of a proteolytically degradable poly(ethylene) glycol scaffold and a non-degradable phosphate and/or polyphosphate nanocomposite hydrogel matrix. This in vitro platform enables quantification of pathogen-induced matrix degradation and screening of the sustained release of phosphate therapeutics in attenuating protease expression. To evaluate matrix degradation as a function of bacterial enzyme levels secreted, we also present a novel method to quantify hydrogel degradation. This method involves staining protease-sensitive hydrogels with Sirius red dye to correlate absorbance of the degraded gel solution with hydrogel weight enabling continuous monitoring of degradation and greater accuracy of degradation kinetics compared to gravimetric measurements. Combined, the proposed in vitro platform and degradation assay provide a novel approach for screening efficacy of therapeutics in attenuating bacterial protease-induced matrix degradation.

Introduction

Severe persistent inflammation due to intestinal disease, including colorectal cancer, ulcerative colitis and Crohn’s disease is potentially fatal if not treated. Common treatments involve antibiotic administration followed by endoscopic or surgical removal and reconnection of diseased portions of the intestinal tract. The process by which successful repair and return of function occur following these insults is complex due to the presence of the intestinal microbiota which, depending on their composition and function, can either enhance or severely impair the healing process. Removal of diseased portions of the intestinal tract leads to the release of pro-inflammatory compensatory host-stress signals which alter the composition and community structure of the normal intestinal microbiome triggering a microbial shift to pathogenic phenotypes.¹ One such phenotype is increased bacterial protease expression resulting in degradation of collagen I and activation of host matrix metalloproteinase (MMP)-9, the latter leading to degradation of collagen IV. This further contributes to increased inflammation, fibrosis, tissue degradation and intestinal wall disruption leading to peritonitis and sepsis. Despite the use of intravenous and oral antibiotics to treat inflammatory bowel disease and to prevent infections during intestinal surgery, patients remain colonized with pathogenic strains following surgical procedures.² This is due in part to the increased prevalence of antibiotic resistance and disruption of normal microbiome community structure. Thus, compounds that suppress pathogen protease production while allowing commensal bacteria to proliferate normally are highly advantageous for prevention and treatment of intestinal tissue degradation without the risk of emergence of resistance. Given the complexity of bacterial-host protease interactions that contribute to intestinal tissue degradation in vivo, the development of in vitro models is highly useful for mechanistic insight on the role of specific host and bacterial enzymes that contribute to this process. In addition, in vitro models also enable systematic screening of novel therapeutics targeting bacterial-host proteolytic activity for prevention of tissue degradation.

Prior studies have demonstrated that surgical injury depletes local intestinal phosphate concentration activating bacterial virulence as microbes acquire sensory systems to detect extracellular phosphate concentration.^2–5 Using both in vitro and in vivo models, we have shown that maintenance of phosphate abundance prevents virulence expression in P. aeruginosa and its transformation to a lethal phenotype.⁴ Our findings also indicate that phosphate and polyphosphate based therapy attenuates in vitro protease production and enhances healing in animal models of intestinal surgical injury and infection, respectively, triggered by Gram-negative and Gram-positive pathogens producing high levels of collagenolytic enzymes.^5,6 Importantly, phosphate and polyphosphate therapeutics result in maintenance of bacterial survival. Given the variation of secreted proteases and phosphate metabolism among microbes as well as the need to maintain intestinal phosphate concentration, we previously developed a drug delivery system to provide sustained release of monophosphate (Pi) and polyphosphate (PPi) combinations from poly(ethylene) glycol (PEG) hydrogel nanoparticles (NP-Pi and NP-PPi, respectively).⁷ Our in vitro findings demonstrated that sustained release of phosphate compounds attenuated in vitro protease production of P. aeruginosa, S. marcescens and E. faecalis.⁷

In the above-mentioned studies, the in vitro efficacy of Pi and/or PPi in attenuating protease activity was evaluated by the addition of bacterial culture media to a commercially available 2D fluorogenic substrate, which is commonly used to quantify cellular enzymatic activity. These substrates contain protease-sensitive peptide sequences derived from collagen which upon proteolytic digestion by gelatinases or collagenases, fluoresce with different intensity proportional to proteolytic activity. This approach for quantifying proteolytic activity, however, does not readily translate to three-dimensional (3D) matrix degradation. Covalently crosslinked synthetic hydrogels are promising biomaterials for therapeutic drug delivery and tissue engineering. Their 3D hydrophilic network is comprised of interstitial crosslinks which allows for their extensive swelling in aqueous solution and biological fluids. Among synthetic hydrogels, those of poly(ethylene) glycol (PEG) provide a variety of advantages for these applications owing to their inherent ease in tuning physical properties (i.e. crosslink density and mesh size) enabling controlled diffusive-based release of therapeutics. Their intrinsic resistance to protein adsorption and cell adhesion, combined with their ability to be readily biochemically modified with key signals of the native extracellular matrix (ECM) render them as a versatile platform for mediating specific cellular functions, including proteolytic degradation.^8–9

In an attempt to analyze the in vitro effectiveness of sustained release of phosphate-based compounds for suppression of proteolytic activity of Gram-negative and Gram-positive pathogens implicated in healing impairment, we developed an in vitro model in which protease producing bacteria were cultured in the presence of a proteolytically degradable crosslinked PEG diacrylate (PEGDA) hydrogel, which served as a degradable tissue surrogate, and a non-degradable PEG hydrogel nanocomposite scaffold encapsulating NP-Pi and/or NP-PPi serving as a Pi and/or PPi releasing reservoir. Although protease-sensitive PEG hydrogels have been extensively used in tissue engineering, the use of crosslinked PEG hydrogel nanoparticles for delivery of hydrophilic therapeutics is far less common. Furthermore, to our knowledge, protease-sensitive PEG hydrogels have not been previously explored as biosensors for detection of proteolytic activity of virulent pathogens or for screening the efficacy of therapeutic compounds that attenuate host-bacterial processes contributing to tissue degradation.

When this biomimetic protease-sensitive scaffold is incubated in the presence of bacterial culture, in the absence of Pi and PPi treatment, it results in complete material degradation with degradation rates specific to each pathogen. This suggests that proteases secreted by each of the three pathogens exhibit different enzymatic activities on matrix degradation. To address the hypothesis that sustained release of phosphates attenuates bacterial protease production leading to matrix degradation, we evaluated the effectiveness of NP-Pi and/or NP-PPi hydrogel nanocomposites in attenuating scaffold degradation by protease producing pathogens in culture. Furthermore, we present a novel method, alternative to gravimetric swollen hydrogel weight measurements, to more accurately quantify degradation kinetics of protease-sensitive hydrogels exposed to enzyme solutions.

Degradation rates of protease-sensitive PEG scaffolds are commonly quantified based on dynamic variations in swollen hydrogel weight during in vitro protease incubation.¹⁰ This method, however, results in inaccurate measurements of gel degradation kinetics, particularly with significant increases in swelling due to network cleavage, and in time estimates for complete material degradation. In this study, matrix degradation is correlated with bacterial enzyme secreted in the medium by staining protease-sensitive PEG hydrogels with Sirius red. Sirius red dye is commonly used for staining collagen by reacting its sulfonic acid groups with basic groups present in the collagen molecule.¹¹ Our theory is that sulfonic groups replace the hydroxyl groups present in the PEG hydrogel using a similar mechanism as in collagen. Upon incubation of protease-sensitive PEG scaffolds with Sirius red, followed by subsequent induction of complete material degradation in protease solution, the absorbance of the solution is correlated with the initial scaffold weight. Stained scaffolds exposed to test pathogens were removed from bacterial cultures co-incubated with the nanocomposite hydrogels and the percentage of gel degradation was quantified via absorbance and correlated with bacterial enzyme levels secreted in the medium. Finally, since pathogens secrete a mixture of proteases which differentially impact extracellular matrix degradation and activation of inactive pro-forms of host MMPs, the efficacy of our proposed degradation assay was evaluated by quantifying gel degradation rates in solutions containing key proteases secreted and purified from P. aeruginosa, including elastase B (LasB) and protease LasA, as well as active human MMP-9.

Materials and methods

Chemicals

Acryl-PEG-SVA (MW= 5,000 g/mol) was purchased from Laysan Bio (Arab, AL). PEGDA (MW= 575 g/mol), Vinyl-2-pyrrolidone (NVP), potassium persulfate (KPS), sodium hexametaphosphate crystalline, Span® 80, Tween® 20, Eosin Y, triethanolamine (TEA), fluorescein isothiocyanate (FITC), 2,2′-(Ethylenedioxy)bis(ethylamine) (EDEA) and piperidine were purchased from Sigma Aldrich (St Louis, MO). Cyclohexane 99%, potassium phosphate monobasic crystals and 2,2’-Azobis(2-methylpropionamidine) dihydrochloride (V-50) were purchased from Acros Organics. Acetone, 1-Ethyl-3-(3-Dimethylaminopropyl)carbodiimide (EDAC), triisopropylsilane, diethyl ether and N,N-diisopropylethylamine (DIEA), were purchased from Fisher Scientific (Hanover Park, IL). All amino acids, resin, O-benzotriazole-N,N,N’,N’-tetramethyluronium-hexafluoro-phosphate (HBTU), N,N-dimethylformamide (DMF) and trifluoroacetic acid (TFA) were purchased from AAPPTec (Louisville, KY). Please add Pi and PPi names and vendor information here. Ultra-high purity (UHP) nitrogen was purchased from Airgas (Radnor, PA, USA). All the other reagents were used without further processing.

Nanoparticle synthesis and purification

Three types of poly(ethylene) glycol diacrylate (PEGDA) nanoparticles were synthesized and used in this study: sodium hexametaphosphate (PPi)-loaded nanoparticles (NP-PPi), sodium monophosphate (Pi)- loaded NPs (NP-Pi) and blank NPs free of phosphates (NP). Nanoparticles were synthesized via inverse miniemulsion polymerization to encapsulate Pi and PPi as previously described.^12,13 Briefly, emulsion consisted of an organic phase of SPAN 80 and 250 mL of cyclohexane and an aqueous PEG precursor solution. The aqueous precursor was comprised of Pi or PPi, the crosslinking agent (PEGDA), the co-monomer NVP, a water-soluble thermal initiator (KPS or V-50), the aqueous phase surfactant Tween 20 and DI water to a final volume of 16.5 mL. All concentrations for aqueous precursor components are indicated in Table 1.

Table 1.

Precursor formulation used for production of phosphate (Pi)- and polyphosphate (PPi)- loaded nanoparticles.

Reagents	Phosphate-loaded NPs (NP-Pi)	Polyphosphate-loaded NPs (NP-PPi)
PEGDA	649 mM	282 mM
NVP	72 mM	121 mM
Salt	85.5 mM	50 mM
Tween 20	478 mg	1611 mg
Initiator	19.28 mM (KPS)	3.42 mM (V-50)
SPAN	8 g	6.87 mg

The aqueous phase was emulsified in the organic phase by first applying a mechanical homogenization using a 10 mm saw-tooth generator (IPRO 250, Pro-Scientific) for 15 min followed by sonication using an ultrasonic horn (SONICS Vibracell VCX750) at a 90% amplitude for 5 pulses of 1 min. During both processes the emulsion was kept on an ice bath. The stable emulsion was transferred to a stirred glass batch reactor and bubbled with nitrogen for 45 min to prevent oxygen inhibition. The reactor was partially immersed in an oil bath allowing a constant reaction temperature of 64°C. The reaction proceeded for 2 hours under a nitrogen blanket. Polymerized nanoparticles were allowed to precipitate overnight at 4 °C after the addition of 100 mL of acetone. Supernatant was removed the next day following the addition of 150 mL of fresh acetone after which precipitated nanoparticles were resuspended by sonication. The new suspension was centrifuged at 3220 g and 4°C for 90 minutes. Following centrifugation and supernatant removal, this cycle was repeated and purified nanoparticles were dried under vacuum for 48 hours. Dried nanoparticle agglomerate was then crushed into a fine powder with a mortar and pestle and stored at room temperature. Blank nanoparticles, used as controls in all experiments, were made using a similar formulation as NP-Pi followed by excessive rinsing for removal and release of Pi from the hydrogel nanoparticle matrix.

Fluorescent labelling of polyphosphate (PPi)

To quantify the amount of polyphosphate (PPi) encapsulated into and released from the nanoparticles, sodium hexametaphosphate (PPi) was conjugated with fluorescein isothiocyanate (FITC). The reaction consisted of a two-step process in which PPi is labeled with amine groups prior to FITC conjugation in order to prevent undesired side reactions.¹⁴ Briefly, 50 mM of PPi in 1M MES buffer was mixed with EDEA (20 equivalents) and EDAC (20 equivalents) followed by pH adjustment to 6.5 with 1M HCl. The mixture was heated for 2 hours at 60°C followed by pH adjustment to 10 with 1M NaOH. The final product from this first step was obtained by precipitation with 45 mL ethanol and centrifugation for 6 min at 1,100g. Precipitation was repeated twice and the final product (PPi-NH₂) lyophilized.

Subsequently, 230 mM of dried PPi-NH₂ in 0.1M carbonate buffer (pH 9) followed by the addition of FITC (4 equivalents). The mixture was stirred at room temperature for 18 hours and purified by precipitation in 45mL ethanol followed by centrifugation for 15 min at 1,100g. Precipitation was repeated twice and the final conjugated FITC-PPi product lyophilized. Fluorescently labeled PPi (FITC-PPi) was then encapsulated into the nanoparticles yielding FITC-PPi loaded nanoparticles (NP-FITC-PPi) based on the protocol described in the previous section.

Nanoparticle characterization

Particle size distribution was quantified using 0.1 mg/mL of nanoparticles in DI water on a nanosizer (NanoSight LM10, Malvern, UK) equipped with nanoparticle tracking analysis software (NanoTracking v3.0). Zeta potential was measured using 0.1mg/mL of nanoparticles in 10 mM NaCl solution on a zetasizer (Nano ZS, Malvern, UK). The mass swelling ratio of nanoparticles, which is inversely correlated with crosslink density and dictates diffusive-based release, was determined gravimetrically as the ratio of swollen to dry nanoparticle weights, M_S and M_D, respectively. The mass swelling ratio was converted to the volumetric swelling ratio (Q) calculated using the following equation:

$$Q= \frac{V_S}{V_D}$$ eq. 1

Where V_s and V_D represent the volume of swollen and dry gel states, respectively obtained by dividing the mass in the swollen state (M_S) by the solvent density (water) and the mass of the dry state (M_D) by the density of PEG (1.12 g cm⁻³).

Hydrogel mesh size estimation

The mesh sizes (ξ) of hydrogel nanoparticles and scaffolds were calculated using the volumetric swelling ratio (eq. 1) measurements based on the Flory-Rehner rubber elasticity theory¹⁵.

$$\xi =V_{2,s}^{-\frac{1}{3}}{\left({\overline{r}}_o{}^2\right)}^{\frac{1}{2}}$$ eq. 2

In equation 2 V_2,S is the polymer volume fraction in the swollen state which is equal to the reciprocal of the volumetric swelling ratio (Q):

$$V_{2,S}= \frac{1}{Q}$$ eq. 3

Where ${\left(\overline{r_o^2}\right)}^{1/2}$ represents the root mean square end-to-end distance between two crosslinks and is described by:

$${\left({\overline{r}}_o{}^2\right)}^{\frac{1}{2}}=l{C}_n^{\frac{1}{2}}{\left(\frac{2\overline{M_c}}{M_r}\right)}^{\frac{1}{2}}$$ eq. 4

In equation 4 l is the average bond length between C-C and C-O in PEG (l=1.46Å) , C_n is the characteristic ratio of PEG (C_n=4), M_r is the molecular weight of PEG repeat unit (M_r=44g/mol), and $\overline{M_c}$ is the average molecular weight between crosslinks, which can be calculated from equation 5:

$$\frac{1}{\overline{M_C}}=\frac{2}{\overline{M_n}}-\frac{\left(\frac{\overline{v}}{V_1}\right)\left[\text{ln}\left(1-V_{2,S}\right)+V_{2,S}+{\chi }_1{V}_{2,S}{}^2\right]}{\left[V_{2,S}{}^{\frac{1}{3}}-\frac{V_{2,S}{}^{ }}{2}\right]}$$ eq. 5

In eq. 5, $\overline{M_n}$ represents the number average molecular weight of the PEG macromer prior to crosslinking, $\overline{v}$ is the specific volume of the polymer (0.84 cm³ g⁻¹), V₁ is the molar volume of the swelling agent (18 cm³ mol⁻¹ for water), χ₁ is the Flory–Huggins polymer solvent interaction parameter (χ₁ = 0.426). All measurements involving nanoparticle characterization were performed on at least three independent samples.

Peptide synthesis and purification

The MMP-sensitive peptide sequence GGVPMS↓MRGGK (↓denotes cleavage point) was synthesized using a Focus Xi peptide synthesizer (AAPPTec, Louisville, KY) by solid-phase peptide synthesis using standard FMOC chemistry, as previously described.¹⁵ Briefly, amino acids were first dissolved in N-methyl-2-pyrrolidinone (NMP) and coupled with N,N-diisopropylethylamine (DIEA) and O-benzotriazole-N,N,N’,N’-tetramethyluronium-hexafluoro-phosphate (HBTU). FMOC groups were deprotected with 20% piperidine in N,N-dimethylformamide (DMF). Peptides were separated from the resin with trifluoroacetic acid (TFA): triisopropylsilane : ddH2O (95:2.5:2.5). Peptide was washed and precipitated in cold diethyl ether and the dried overnight products dissolved in ddH2O and purified by reverse-phase high-performance liquid chromatography (HPLC) (Agilent 1200 series system). The final products were lyophilized and stored at −80°C.

Synthesis of PEGDA-peptide crosslinker

A photopolymerizable macromer of PEGDA containing an MMP-sensitive peptide between its terminal acrylate groups was synthesized and used to create a proteolytically degradable hydrogel scaffold.¹⁵ Briefly, the PEDGA-peptide crosslinker was synthesized by conjugating Acryl-PEG5000-SVA with MMP-sensitive peptide in a 2:1 molar ratio. The reaction was carried out in the dark in 50 mM NaHCO₃ solution (pH 8) for 4 h. This reaction resulted in the synthesis of the protease-sensitive PEGDA crosslinking macromer, Acryl-PEG5000- GGVPMS↓MRGGK-PEG500-Acryl (MW ~ 11,000 Da). Following conjugation, the macromer was dialysed for 24 h, lyophilized and stored at −20°C until use.

Sirius red method for quantification of hydrogel degradation

Synthesis and staining of protease-sensitive PEG hydrogel.

PEGDA hydrogels, functionalized with the proteolytically degradable peptide sequence GGVPMS↓MRGGK, were used as a degradable tissue surrogate. Hydrogel precursor was prepared with 1x HEPES buffered saline (HBS) (pH 7.4), 2 mM of the protease PEGDA crosslinker (MW ~ 11,000 Da), 37 mM NVP, 225 mM of co-initiator TEA and 0.05 mM photosensitive dye Eosin Y. Precursor’s pH was adjusted to 7.4 by addition of HCl. A volume of 100 μL of precursor solution was added into as well of a 96-well plate followed by photopolymerization. Hydrogels were polymerized for 5 minutes using visible light (λ = 514nm) from an Argon Ion Laser (Coherent, Inc., Santa Clara, CA) and laser flux of 100 mW cm⁻². Each polymerized gel was incubated for 24 hours in 500μL of Sirius red (Direct Red 80, Sigma-Aldrich). Subsequently, degradable gels were removed from the Sirius red solution and incubated in HBS for an additional 24 hours. Cell culture and degradation experiments (described below) were performed after gels achieved equilibrium swelling following incubation for 48 hours in aqueous solution.

Standard curve for quantification of hydrogel degradation.

After staining degradable hydrogels with Sirius red dye, gels were removed from solution and incubated in HBS solution to ensure that the dye was fully rinsed prior to quantification of gel degradation using absorbance measurements. No changes in the absorbance of the medium surrounding fully rinsed Sirius red stained hydrogels were noted for several days (data not shown). This confirmed that Sirius red staining of hydrogels does not result in release of the dye by diffusion and that the measured changes in absorbance are due to hydrogel degradation.

Sirius red stained hydrogels were sectioned and weighed in centrifuge tubes with gel weights ranging from 25 to 120 mg. A volume of 1 mL of 10 μg/mL collagenase standard from Clostridium histolyticum (Sigma-Aldrich) was added in each tube. After full hydrogel degradation in collagenase solution was achieved, a volume of 200μL of the degraded hydrogel solution was added to a 96-well plate and absorbance was measured at 540 nm. The absorbance of the collagenase standard solution was also measured and subtracted from all hydrogel absorbances to account for possible background. A standard curve was used to correlate hydrogel initial weight and absorbance measured after degradation. Non-degradable PEG hydrogels were also stained with Sirius red and used as negative controls. No variations or increases in absorbance over time were noted in the case of non-degradable gels incubated with collagenase using the Sirius red protocol indicating that the assay can be accurately used for quantification of gel degradation (data not shown).

Synthesis of non-degradable nanocomposite PEG hydrogels

PEDGA nanocomposite hydrogels were used as a reservoir for sustained release of phosphates and polymerized using free-radical photopolymerization as described previously.¹⁶ Briefly, non-degradable nanocomposite hydrogel precursor was prepared with 1x bacterial growth tryptone yeast (TY) media, 1.5 mM PEGDA crosslinker (MW 12,000), 37 mM NVP, 225 mM of co-initiator TEA and 0.05 mM photosensitive dye Eosin Y. Dried nanoparticles were weighted and added to a 96-well plate following by the addition of 100 μL hydrogel precursor. The mixture was photopolymerized using an Argon Ion Laser at 514 nm and laser flux of 100 mW cm⁻² for 5 min. Encapsulated blank nanoparticles were pre-incubated in TY media prior to experiments, to avoid possible uptake of nutrients from the medium which would induce decreases in bacterial growth (data not shown).

Polyphosphate (PPi) release kinetics from nanoparticles and nanocomposite scaffolds

Release kinetics of FITC-PPi from nanoparticles and nanocomposite scaffolds were quantified by measuring the fluorescence of released samples using a microplate reader (SpectraMax M2, Molecular Devices). In the case of measurements of PPi release from free nanoparticles, 1% (w/v) FITC-PPI-NPs were added directly with DI water to a microcentrifuge tube. At pre-determined time points tubes were centrifuged, supernatant removed, and fresh DI water added. A volume of 200 μL of the removed supernatant was added to a 96-well plate and fluorescence was measured using a plate reader. To quantify release of PPi from nondegradable nanocomposite scaffolds, the nanocomposite was placed in a 24 well-plate following the addition of 1ml of DI water. At pre-determined time points the media was removed, fluorescence measured, and fresh media added to each well. Fluorescence measurements were adjusted to account for photobleaching effects of FITC-PPi nanoparticles upon exposure to visible light photopolymerization to form the nanocomposites (data not shown). In both cases fluorescence measurements were converted into cumulative concentration, using a standard curve to quantify FITC-PPi release kinetics.

Collagenolytic bacterial strains

Gram-positive (E. faecalis) and Gram-negative (P. aeruginosa and S. marcescens) pathogens expressing high proteolytic activity were used in all experiments.^5,17–18 P. aeruginosa PAO1 derivative XEN41 bioluminescent strain was purchased from Calpier Life Sciences, Inc. S. marcescens ICU2–4lum strain was isolated from an ICU patient at the University of Chicago and further transformed into bioluminescent according to our previously published protocol using a pAKlux2 plasmid containing the luxCDABE luciferase cassette (Addgene, Ref #14080).^19,20 E. faecalis E61 was isolated from rat anastomotic tissue at the University of Chicago. Bacterial strains from frozen stocks were plated in tryptone yeast (TY) plates and incubated overnight at 37°C. An inoculation loop of plated bacteria was cultured overnight at 37°C in 2mL of TY media. Liquid cultures were diluted 1:100 in fresh TY media prior to all experiments.

Bacterial growth in the presence of hydrogel nanocomposites

Bacterial growth (OD) was evaluated for each strain in the presence and absence of non-degradable hydrogel nanocomposite scaffolds. The groups tested in vitro for this purpose included a control group (no hydrogel in culture), a nanocomposite hydrogel matrix loaded with 1% (w/v) blank nanoparticles (NP) pre-incubated in TY media and nanocomposite hydrogel matrices loaded with either 1% (w/v) NP-Pi and 1% (w/v) NP-PPi. Hydrogel matrices were added to a 24 well-plate and incubated with 1 mL of 1:100 dilution bacterial culture at 37°C.

Efficacy of sustained release of phosphates in attenuating hydrogel degradation by pathogen secreted proteases

To screen the efficacy of sustained release of Pi and PPi in attenuating bacterial protease-induced matrix degradation, a physiologically relevant in vitro assay was developed. Phosphate (-Pi) and polyphosphate (-PPi) nanoparticles (NP-Pi and NP-PPi, respectively), as well as blank nanoparticle controls (NP), were encapsulated in non-degradable hydrogels using photopolymerization to form hydrogel nanocomposites. Non-degradable nanocomposite matrices of different types (NP, NP-Pi, NP-PPi and combinations of NP-Pi and NP-PPi), were co-incubated separately with the proteolytically degradable stained hydrogel matrix in a 24-well plate (Fig.1). All groups are summarized in Table 2.

fig. 1.

schematic of in vitro assay for screening bacterial proteolytic activity in the presence and absence of np-pi and/or np-ppi treatment. co-incubation of a sirius red stained proteolytically degradable hydrogel scaffold and a non-degradable phosphate (pi) and/or polyphosphate (ppi) releasing non-degradable nanocomposite hydrogel in bacterial culture.

Table 2.

Summary of experimental groups used to screen pathogen induced protease-degradation and attenuation in vitro.

Groups	Components
Control 1 (no gel)	Bacterial culture
Control 2	Bacterial culture + Sirius red stained degradable hydrogel
NP	Bacteria culture + Sirius red stained degradable hydrogel + (Blank NP) nanocomposite hydrogel
NP-Pi	Bacteria culture + Sirius red stained degradable hydrogel + (NP-Pi) nanocomposite hydrogel
NP-PPi	Bacteria culture + Sirius red stained degradable hydrogel + (NP-PPi) nanocomposite hydrogel
NP-PPi + NP-Pi	Bacteria culture + Sirius red stained degradable hydrogel + (NP-PPi + NP-Pi) nanocomposite hydrogel

Overnight growth of bacterial strains was diluted 1:100 in TY media and a volume of 1mL of each dilution was added separately to each well. Two control groups were used to quantify protease-induced matrix degradation and attenuation based on Pi and PPi treatment in culture. The first control consisted of bacterial incubation in the presence of the Sirius red stained degradable hydrogel scaffold in the absence of Pi or PPi nanocomposite treatment. The second control group included bacterial culture alone in the absence of gel (nanocomposite or the proteolytically degradable scaffold). The incubation time required for each pathogen was based on the time at which respective hydrogel treatment groups from controls were fully degraded. When this was achieved, the optical density (OD 600nm) was measured to quantify bacterial growth, and the remaining gels from each group were removed from the wells and added separately to 1.5 mL centrifuge tubes after which remaining weight was measured by both swollen hydrogel weight gravimetric measurements and as a function of absorbance. To measure absorbance, 1 mL of 10 μg/mL collagenase solution was added to the centrifuge tube until complete degradation of the remaining protease-sensitive hydrogel was achieved. The tubes were centrifuged for 5 min at 12000 rpm, 200μL of fully degraded solution was added to 96-well plate and absorbance was measured at 540 nm. The standard curve of weight and absorbance allowed for accurate quantification of degradation percentage for each treatment group. The percentage of degradation was correlated to the concentration of bacterial enzyme secreted in the medium. Normalized enzyme levels were obtained by normalizing percent degradation to the OD for each sample.

Overproduction and purification of P. aeruginosa proteases

LasA and LasB, were amplified from Pseudomonas aeruginosa Xen41 chromosomal DNA as a template and inserted into pJL1 vector with 6 His-tag. E. coli BL21 (DE3) Star cells were transformed with the constructed plasmid pJL1-LasA and LasB. To produce protein, an overnight culture of BL21(DE3) Star harboring plasmid was diluted in 250 mL of LB with 50μg/mL of kanamycin in the flask and incubated at 37 °C at 220 rpm until OD600nm of ~0.5. Then, isopropyl β-D-1-thiogalactopyranoside (IPTG) was added to induce enzyme production and incubated at 18 °C at 220 rpm overnight. Cells were harvested by centrifugation, resuspended with lysis buffer and lysed by sonication. The enzyme was purified through the 6 His-tag column and concentrated by ultrafiltration concentrators. The size of protein was confirmed by loading on SDS-PAGE, and the concentration was measured by a Quick-Start Bradford protein assay kit.

Quantification of matrix degradation kinetics

Sirius red stained protease-sensitive PEG scaffolds were used for evaluating gel degradation kinetics in the presence of activated human MMP-9 (Sigma-Aldrich), and P. aeruginosa extracted LasA and LasB. Hydrogels were incubated separately in centrifuge tubes containing 1 mL of HBS media and 5 μg/ml of either LasA, LasB or active MMP-9. Absorbance was measured from time zero until full gel degradation. At each time point, an aliquot of 200 μL of the surrounding solution was added to a 96-well plate and absorbance was read at 540 nm. The aliquoted solution used was returned to the centrifuge tube at each time point to quantify cumulative degradation over time. The absorbance at each time point was converted to degradation percentage for all groups.

Statistical analyses

All experiments were performed in triplicate and quantitative results were presented as the mean ± standard deviation. Analysis of variance (ANOVA) of experimental groups compared with controls was used and statistical significance was considered for p < 0.05, unless otherwise noted.

Results and discussion

Nanoparticle characterization

The different types of nanoparticles (NP-Pi, NP-PPi and NP-FITC-PPi) were characterized in terms of their swelling ratio, particle size distribution and zeta- potential. Nanoparticle swelling ratio is an important hydrogel property for characterizing diffusive-based drug delivery since it is inversely related to network crosslink density and dictates drug release kinetics. The swelling ratios of NP-PPi and NP-FITC-PPi were found to be similar (5.59 ± 0.06 and 5.75 ± 0.28, respectively) as expected since they were polymerized under identical formulations and conditions. The swelling ratio of NP-Pi was found to be lower (4.8 ± 0.09) compared to NP-PPi and NP-FITC-PPi. This is attributed to differences in precursor formulation used to produce nanoparticles of lower crosslink density to sustain release of Pi due to its inherent increased diffusivity in gels of similar network properties compared to PPi. The zeta-potential, a measure of particle surface charge characteristics, was found to be −9.37 mV ± 2.53 mV and −20.34 mV ± 0.79 mV for NP-Pi and NP-FITC-PPi, respectively. An increase in negative surface charge of NP-FITC-PPi is most likely attributed to the encapsulated FITC conjugated compound. Finally, the zeta-potential of NP-Pi was −17.92 mV ± 1.05 mV, which was more negative relative to NP-PPi. This was expected due to the potential increased diffusivity of Pi within the crosslinked network and its accumulation near the nanoparticle surface. In terms of particle size, the diameters of NP-Pi and NP-FITC-PPi were similar with slight decreases in particle size noted for NP-PPi. All values of nanoparticle properties are reported in Table 3.

Table 3.

Nanoparticle characterization.

	Nanoparticle Diameter	Swelling Ratio	Zeta Potential (mV)
NP-Pi	181 ± 57	4.8 ± 0.09	−17.92 ± 1.05
NP-PPi	147 ± 57.3	5.59 ± 0.06	−9.37 ± 2.53
NP-FITC-PPi	185.1 ± 83.4	5.75 ±0.28	−20.34 ± 0.79

The Flory-Rehner theory was used to estimate the mesh size of both NP-PPi and NP-FITC-PPi from swelling measurements (Table 3). Similar to swelling ratio findings, no statistical significant differences in mesh size between NP-Pi and NP-FITC-PPi were noted, suggesting that encapsulation of FITC-PPi in the nanoparticles does not impact nanoparticle network properties or diffusive characteristics. (Fig. 2).

fig. 2.

mesh size estimation for np-ppi and np-fitc-ppi. values reported in terms of mean ±standard deviation (n = 3).

Quantification of nanoparticle release kinetics

Drug release from polymeric nanoparticles usually involves an initial burst due to molecule accumulation at the particle surface, followed by slow and controlled release.²¹ In order to obtain measurements of PPi release kinetics, sodium hexametaphosphate (PPi) was fluorescently labeled with fluorescein isothiocyanate (FITC). The labeling procedure reproduced end-modified PPi (FITC-PPi) in which one FITC molecule is incorporated at each end of PPi molecule.¹⁴ Therefore, in the most likely scenario of potential hexamer dissociation, fluorescence would not change and PPi concentration could be measured in a more stable and reproducible manner.

To measure the concentration of FITC-PPi being released from nanoparticles over time, a standard curve was created correlating fluorescence with concentrations ranging from 0.1 to 1 mg/ml. The resulting curve, provided as supplementary data (Fig. S1), demonstrated linear behavior with a coefficient of determination of 0.9971. The slope and y-intercept were used to further convert the fluorescence values obtained from the release kinetics into PPi concentration. Cumulative release kinetics of PPi from nanoparticles was performed at 37°C under perfect sink conditions. Release kinetics of FITC-PPi from free NP-PPi indicate a high burst release in the first 2 hours (Figure 3). As shown in Figure 3, release of FITC-PPi from the nanocomposite are delayed compared to its release from free NP-PPi.

fig. 3.

cumulative release kinetics of fitc-ppi, from free nanoparticles and the nanocomposite over 24 hours. values reported in terms of mean ± standard deviation (n = 3).

Separate in vitro experiments performed with each test pathogen required different incubation times, all of which did not exceed 24 hrs. Therefore, using the standard curve (presented in Fig. S1), the equivalent PPi concentration released up to 24 hours is estimated to be 0.31 mg/ml and 0.14mg/ml for the nanoparticles and nanocomposite, respectively.

Sirius red staining of Protease-Sensitive PEG Scaffolds for Quantifying Hydrogel Degradation Kinetics

The most commonly used method to measure hydrogel degradation is based on dynamic gravimetric measurements in hydrogel swollen weight during enzyme incubation. This method, however, makes it difficult to accurately quantify gel degradation kinetics as the network starts to degrade and as the hydrogel weight increases due to increases in swelling. Furthermore, precise quantification of the time required to achieve complete material degradation becomes difficult to capture. To more accurately measure hydrogel matrix degradation kinetics as well as the time required for complete material degradation, we developed a new method whereby a degradable PEG hydrogel scaffold is stained with Sirius red after polymerization (Fig. 4), and its weight is measured as a function of absorbance (540 nm) upon full degradation. Using this method, we hypothesize that PEG hydrogel staining occurs through a mechanism similar to collagen, in which sulphonic groups from Sirius red react with basic groups (e.g. hydroxyls) from PEG chains.

fig. 4.

images of a degradable peg scaffold hydrogel immediately following polymerization (left) and after staining with sirius red (right).

After scaffolds were stained with Sirius red, a standard curve of scaffold weight as a function of absorbance was created. Sectioned pieces of the stained hydrogel, ranging from 12 mg to 127 mg, were first fully degraded in a solution containing 10 μg/mL of collagenase from Clostridium histolyticum (Sigma-Aldrich). After full gel degradation was achieved, the absorbance of the final solution was measured and correlated with initial gel weight (Figure S2). The relationship between gel weight and absorbance was found to be linear (r²=0.9994), which enabled quantification of hydrogel matrix degradation in a less labor intensive and more reliable manner as indicated below. It is important to highlight that the enzymatic activity of the collagenase used does not affect the relation between weight and absorbance. This means that the same standard curve would be obtained if a different concentration or a different enzyme capable of degrading a hydrogel scaffold of the same material properties and biochemical composition is utilized.

Bacterial growth in the presence of hydrogel nanocomposite

To verify that the presence of the hydrogel nanocomposite scaffold treatment in bacterial culture did not impact bacterial growth, different hydrogel nanocomposites were added to a bacterial culture and incubated in the presence of bacteria and growth (OD) was quantified. The control group, without treatment, was used as a standard for comparisons of treatment groups in running statistical analyses. The treatment groups for all three pathogens included non-degradable nanocomposites encapsulated with 1% NP-PPi, 1% NP-Pi or a mixture of 1% NP-PPi + 1% NP-Pi. Blank nanoparticles at 1% and 2% (w/vol) were also encapsulated into the hydrogels yielding a negative nanocomposite control. The data in Fig. 5 indicate no statistical differences in bacterial growth for all test pathogens cultured in the presence of non-degradable nanocomposite hydrogels.

fig. 5.

optical density (od) of (a) s. marcescens, (b) p aeruginosa and (c) e. faecalis in the presence of non-degradable pi and or ppi nanocomposite peg hydrogels. values reported in terms of mean ±standard deviation (n = 3).

Hydrogel degradation by Pathogen Secreted Proteases

PEG hydrogels are widely used as biomimetic matrices, due to their biocompatibility, hydrophilicity and precise control over physical and chemical properties.¹⁵ Photopolymerization of PEG crosslinkers functionalized with the protease-sensitive peptide sequence (GGVPMS↓MRGGK) results in the formation of hydrogels with susceptibility to degradation by human and bacterial proteases. The incorporation of this sequence induces dynamic changes in hydrogel physical and mechanical properties in enzyme incubation as shown in our recently published study.²² The initial elastic modulus (prior to enzyme incubation) of the protease-sensitive scaffold used in the present study, was approximately 2 kPa based on the precursor composition specified.²² The proteolytic activity of P. aeruginosa, S. marcescens and E. faecalis based on phosphate treatment was measured in the presence of the degradable scaffold and the Pi and PPi nanocomposite reservoir in culture (Fig. 6).

fig. 6.

in vitro protease activity assay in vitro set-up. bacterial culture is carried out in the presence of a proteolytically degradable peg hydrogel scaffold stained with sirius red (enabling monitoring and quantification of matrix degradation by proteases produced by pathogens) and a non-degradable phosphate (pi) and/or polyphosphate (ppi) nanocomposite peg hydrogel serving as a reservoir of sustained release of pi and/or ppi for attenuation of protease production.

The mesh sizes of the non-degradable and degradable scaffolds were calculated using gravimetric measurements and further Flory-Rehner rubber elasticity theory. The estimated mesh size of both hydrogel scaffolds is between 16–18 nm, which is lower than the nanoparticle diameter and bacteria dimensions (Fig. 7). This suggests that the nanoparticles are confined inside the nanocomposite hydrogel and that bacteria do not migrate inside the scaffolds.

fig. 7.

estimated mesh sizes of nanocomposite and degradable hydrogel scaffolds. values reported in terms of mean ±standard deviation (n = 4).

An increase in bacterial growth is noted for the treatment groups as compared to control for S. marcescens and E. faecalis (Fig. 8A, C). The specific reason for this increase is unknown as no differences in growth were observed between the two control groups. These findings suggest that the degradation products of the stained degradable scaffold in the media and the sustained release of Pi and PPi from the nanocomposite reservoirs do not attenuate bacterial growth (Fig. 8). The noted increases in growth did not impact Pi or PPi treatment groups since NP-Pi and NP-PPi nanocomposite groups result in the maintenance of bacterial survival and reduction in E. faecalis and S. marcescens induced proteolytic scaffold degradation levels, respectively. In the case of P. aeruginosa growth, no statistical significant changes among treatment groups and controls were noted (Fig. 8E).

fig. 8.

bacterial protease activity quantified as a function of hydrogel degradation percentage (d, e, f) and growth in terms of od (a, b, c) in the presence and absence of phosphate-based treatment. growth and percent matrix degradation for s. marcescens (a, d), p. aeruginosa (b, e), and e. faecalis (c, f). data reported in terms of mean ± standard deviation (n = 4). (*, +) indicate statistical significance (p<0.05) compared to control 1 (no gel) and control 2, respectively.

As mentioned previously, the incubation time for each pathogen was defined as the time at which complete degradation of hydrogel control groups was achieved. After this time point, stained hydrogels that only partially degraded were removed from culture for further analysis of degradation percentage. Treatment groups consisting of 1% NP-PPi and 1% NP-Pi + 1% NP-PPi in nanocomposite gels were effective in attenuating matrix degradation induced by both Gram-negative pathogens but were ineffective for Gram-positive E. faecalis. In contrast, the 1% NP-Pi nanocomposite group attenuated E. faecalis induced matrix degradation, yet this treatment was less effective and ineffective in suppressing P. aeruginosa and S. marcescens induced-proteolytic degradation, respectively. Although not investigated in this study, further adjustments in the combination formulation (NP-Pi and NP-PPi) may potentially provide broad-spectrum protection against proteolytic activity of both Gram-positive and Gram-negative pathogens.

In the data presented in Figure 8 (D–F), the percentage of gel degradation was quantified based on measurements in gel swollen weight and degraded weight approximated using the proposed Sirius red staining protocol. For S. marcescens and E. faecalis the swollen hydrogel weight of the remaining hydrogel after removal from bacterial culture was found to be greater than the initial weight prior to enzyme exposure, which does not accurately depict actual material degradation. This is attributed to the degradation of crosslinks from the core structure of hydrogel, which allows for further swelling and higher swollen weight after incubation. In the case of P. aeruginosa both methods used to quantify matrix degradation (gravimetric measurements in swollen weight and gel weight measured by Sirius red staining), were able to capture percent hydrogel degradation in bacterial culture. The Sirius red staining protocol, however, appears to be more accurate as it resulted in smaller standard deviations from the mean, strengthening its ability to quantify protease degradation as compared to swollen weight measurements.

Furthermore, hydrogels degraded by P. aeruginosa exhibited significant decreases in diameter over time (data not shown) which did not occur in the case of the other pathogens that demonstrated uniform gel degradation. Since the degradable scaffold serves as a substrate for bacterial enzymes and its degradation is reaction-diffusion mediated, the degradation reaction depends on both Michaelis-Menten kinetics and Fick’s second law of diffusion.^23,24 Therefore, we believe that noted differences in hydrogel behavior observed between the different pathogens may be related to the reaction or diffusion dominance of the degradation process, however, further investigation is required to confirm this hypothesis.

Degradation Kinetics of P. aeruginosa Secreted Proteases

Pathogens secrete a mixture of proteases that may act differently on extracellular matrix degradation as well as activate host proteases, including MMPs which further contribute to tissue breakdown. The most abundant proteases identified to be expressed by S. marcescens and E. faecalis are the zinc metalloprotease SM, and gelatinase (gelE) enzymes, respectively.^25–28 Main proteases secreted by P. aeruginosa, are elastase B (LasB), protease LasA and alkaline protease (AprA).²⁸ To demonstrate the effectiveness of our proposed Sirius red protocol in quantifying pathogen specific hydrogel degradation kinetics, gel degradation rates were quantified in the presence of overproduced and purified P. aeruginosa enzymes, elastase B (LasB) and protease LasA, as well as by human active MMP-9. MMP-9 is a host matrix metalloprotease whose inactive pro-form is activated by bacterial proteases leading to degradation of collagen IV and contributing to impaired intestinal wound healing in animal models of surgical infection and healing, such as the incidence of anastomotic leak.²⁹ For each case mentioned above degradation rates were obtained using the Sirius red protocol were compared to those obtained through gravimetric measurements in swollen gel weight.

Figure 9A displays dynamic changes in absorbance measured at each time point in the absence of enzyme (control) and in the presence of LasA, LasB or active human MMP-9 enzyme solutions. Gel degradation kinetics were then quantified by correlating absorbance to remaining gel weight as shown in Figure 9B. Using the absorbance method, a smooth decrease in degraded gel weight is obtained until complete hydrogel degradation is achieved. The times for complete gel degradation in the presence of LasB and MMP-9 active enzymes were found to be 4.5 and 7.5 hours, respectively, while LasA did not induce matrix degradation. Furthermore, LasB was found to degrade the hydrogel matrix approximately 1.7 times faster than MMP-9. These findings are significant as they provide insight on the role of P. aeruginosa secreted proteases on collagen degradation and its pathogenesis leading to healing impairment. Comparisons of degradation profiles based on gravimetric measurements in swollen gel weight display a noted increase in weight due to increases in hydrogel swelling as a result of matrix degradation, followed by a rapid decrease in weight with further increases in degradation. The ability to accurately capture the time required to achieve complete material degradation is not possible when relying on gravimetric swollen hydrogel measurements in hydrogel weight (Fig. 9C).

fig. 9.

comparison of degradation kinetics of proteolytically degradable hydrogel scaffolds in the presence human mmp-9 and p. aeruginosa expressed las a and las b obtained using the sirius red staining protocol and gravimetric measurements in hydrogel swollen weight. (a) dynamic measurements of sirius red stained hydrogel absorbance and (b) converted weight variations obtained from the absorbance measurements shown in (a) using the standard curve shown in fig. s2. (c) dynamic variations in hydrogel swollen weight obtained gravimetrically. values represent mean ± deviation (n = 3).

Dynamic gravimetric measurements in hydrogel swollen weight do not represent the actual material degradation profile as these cannot discriminate between polymer weight from the aqueous weight imbibed by the swollen hydrogel network. In addition, this approach for quantifying the degradation kinetics of enzymatically degradable hydrogels is more labor-intensive as the enzyme solution needs to be removed and replenished each time the swollen weight is measured during the experiment. The absorbance method allows for continuous monitoring of decreases in gel weight until full degradation is achieved and more accurately captures the time at which complete material degradation occurs in enzyme solution. The 3D in vitro method presented combined with our proposed assay for quantifying proteolytic degradation, allowed for efficient screening of pathogen-induced proteolytic activity and sustained delivery of phosphate-based therapeutics in attenuating bacterial protease-induced matrix degradation. Since PEG hydrogels can be easily modified, they can be engineered to be degradable by specific peptide substrates to mechanistically screen and identify proteases secreted by pathogens that contribute to tissue breakdown and activation of host MMPs that contribute to matrix degradation.

Conclusions

In this work a novel in vitro approach for assessing the impact of proteases released by collagenolytic pathogens on PEG hydrogel degradation was evaluated. In order to accurately quantify hydrogel degradation in the presence of bacteria as well as matrix degradation kinetics as a result of pathogen-secreted key enzymes and host MMPs, a new method to more accurately quantify matrix degradation rates was developed. Staining proteolytically degradable hydrogels with Sirius red was found to provide greater accuracy for quantifying hydrogel degradation kinetics as compared to gravimetric measurements in swollen gel weight. Our proposed in vitro platform also allowed us to screen the efficacy of sustained release of phosphate therapeutics in attenuating pathogen-induced matrix degradation. Sustained release of PPi and a combination treatment of PPi and Pi provided by nanocomposite hydrogels in bacterial culture significantly attenuated Gram-negative induced matrix proteolytic degradation. Conversely, in the case of Gram-positive pathogens, decreases in proteolytic degradation was achieved with sustained release of Pi. Lastly, quantification of degradation matrix degradation kinetics by specific proteases produced by P. aeruginosa as well as human MMP-9 indicated that protease LasA released does not influence matrix degradation, while LasB degrades hydrogel matrix in approximately 1.7 times faster than human MMP-9. Our presented approach holds promise in the design of in vitro studies that may potentially provide mechanistic insight on the microbiology of host-pathogen interactions and identification of therapeutics in attenuating cell-mediated responses that contribute to matrix degradation and impaired wound healing.