Engineered aprotinin for improved stability of fibrin biomaterials

Biomaterials 32 (2011) 430e438

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Biomaterials journal homepage: www.elsevier.com/locate/biomaterials

Engineered aprotinin for improved stability of fibrin biomaterials Kristen M. Lorentz a, Stephan Kontos a, Peter Frey a, b, Jeffrey A. Hubbell a, b, c, * a

Institute of Bioengineering, Ecole Polytechnique Fédérale de Lausanne (EPFL), 1015 Lausanne, Switzerland Department of Pediatric Urology, Centre Hospitalier Universitaire Vaudois (CHUV), 1011 Lausanne, Switzerland c Institute of Chemical Sciences and Engineering, Ecole Polytechnique Fédérale de Lausanne (EPFL), 1015 Lausanne, Switzerland b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 18 August 2010 Accepted 30 August 2010 Available online 22 September 2010

Fibrin has been long used clinically for hemostasis and sealing, yet extension of use in other applications has been limited due to its relatively rapid resorption in vivo, even with addition of aprotinin or other protease inhibitors. We report an engineered aprotinin variant that can be immobilized within fibrin and thus provide extended longevity. When recombinantly fused to a transglutaminase substrate domain from a2-plasmin inhibitor (a2PI1e8), the resulting variant, aprotinin-a2PI1e8, was covalently crosslinked into fibrin matrices during normal thrombin/factor XIIIa-mediated polymerization. Challenge with physiological plasmin concentrations revealed that aprotinin-a2PI1e8-containing matrices retained 78% of their mass after 3 wk, whereas matrices containing wild type (WT) aprotinin degraded completely within 1 wk. Plasmin challenge of commercial sealants Omrixil and Tisseel, supplemented with aprotinin-a2PI1e8 or WT aprotinin, showed extended longevity as well. When seeded with human dermal fibroblasts, aprotinin-a2PI1e8-supplemented matrices supported cell growth for at least 33% longer than those containing WT aprotinin. Subcutaneously implanted matrices containing aprotinin-a2PI1e8 were detectable in mice for more than twice as long as those containing WT aprotinin. We conclude that our engineered recombinant aprotinin variant can confer extended longevity to fibrin matrices more effectively than WT aprotinin in vitro and in vivo. Ó 2010 Elsevier Ltd. All rights reserved.

Keywords: Fibrin Degradation Recombinant protein Crosslinking

1. Introduction Fibrin is an integral component of the biologically active clot within healing wounds and has been extensively characterized as a biomaterial. Fibrin offers numerous advantageous properties, including facile autologous and allogenous isolation, inherent biological activity arising from its integrin and growth factor-binding sites, and natural protease-dependent resorption in vivo. Fibrin sealant has been approved for an array of indications by the United States FDA [1] and other regulatory bodies and continues to be explored clinically as a delivery vehicle for multiform therapeutics, such as small molecule drugs [2], growth factors [3], and stem cells [4]. In addition, many research efforts have focused on utilizing fibrin hydrogels as cell-supporting scaffolds in tissue engineering of adipose, cardiovascular, ocular, muscle, liver, skin, cartilage, and bone tissues [5]. Yet, despite its advantages, fibrin’s applicability can be limited due to its relatively rapid rate of resorption in vivo.

* Corresponding author. EPFL-SV-IBI-LMRP, Station 15, CH-1015 Lausanne, Switzerland. Tel.: þ41 216939681; fax: þ41 216939685. E-mail address: jeffrey.hubbell@epfl.ch (J.A. Hubbell). 0142-9612/$ e see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biomaterials.2010.08.109

Technologies that can reduce the rate of fibrin degradation may thus improve clinically-used fibrin products as well as advance tissue engineering applications as future clinical solutions. Fibrin is sensitive to proteolytic degradation by enzymes, such as matrix metalloproteinases and plasmin [6]. Plasminogen, the precursor of plasmin, is present in most fibrin preparations and is cleaved by plasminogen activators produced by cells in vitro and in vivo to yield the active fibrin-degrading enzyme plasmin. Matrix metalloproteinases are produced or activated by almost all cell types. Strategies have been explored to prolong the life of fibrin matrices, ranging from reinforcement with synthetic polymers, such as polyethylene glycol [7], to addition of protease inhibitors [8e10]. One such inhibitor, aprotinin (bovine pancreatic trypsin inhibitor, Bayer’s TrasylolÒ), is a small (6.5 kDa), monomeric serine protease inhibitor found to effectively inhibit the activity of several proteases, including plasmin, trypsin, chymotrypsin, and kallikrein [11]. Yet, although aprotinin is an efficient inhibitor of fibrin degradation, it rapidly diffuses out of fibrin matrices and thus does not provide extended matrix protection outside of an in vitro setting in which its escape is contained. Recently, a method has been devised to prevent this diffusive escape by chemically conjugating aprotinin to fibrinogen prior to gel formation in an

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organic phase reaction scheme; these authors showed prolonged presence of fibrin to at least 4 d in a chick chorioallantoic membrane invasion assay [12]. We sought to develop an aprotinin variant that could be covalently bound to fibrin during gel formation and thus confer extended longevity to fibrin matrices, especially in vivo. Previously, our laboratory has developed a method to covalently bind bioactive factors to fibrin matrices via exploitation of an octapeptide transglutaminase substrate sequence taken from a2-plasmin inhibitor, which is naturally present at high densities within endogenous fibrin clots [13]. By recombinantly fusing the a2PI1e8 substrate sequence tag to aprotinin, the resulting purified variant, aprotinin-a2PI1e8, could be covalently crosslinked into fibrin matrices during normal thrombin and factor XIIIa-mediated polymerization. Such a method would be advantageous in that aprotinin-a2PI1e8 is simple to express and purify using standard techniques and that the mutant inhibitor would be crosslinked to sites on fibrin which correspond to the natural sites of the physiologically native plasmin inhibitor, a2-plasmin inhibitor. By comparing aprotinin-a2PI1e8 with wild type (WT) aprotinin in in vitro and in vivo settings, we demonstrate that our aprotinin variant can prolong the stability of fibrin matrices more effectively than the WT form of the protein.

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2.5. Fluorescent fibrinogen derivatives For in vitro plasmin challenges, fluorescein conjugation to fibrinogen was performed by reacting 2 mg of fibrinogen with 20 equivalents (0.116 mmol) fluorescein isothiocyanate (FITC, Sigma-Aldrich) at room temperature in PBS at pH 7.4 overnight. FITC-conjugated fibrinogen was purified from the reaction mixture by size exclusion chromatography on a 12 mL Sephadex G-50 column (Sigma-Aldrich), while monitoring the fluorescence of elution fractions on a UV lamp. For in vivo fibrin gel studies, AlexaFluor-680 modification of fibrinogen was performed by reacting 1 mg of AlexaFluor-680 carboxylic acid succinimidyl ester (Invitrogen), dissolved in dimethyl sulfoxide, with 10 mg of fibrinogen in 100 mM sodium bicarbonate, 150 mM NaCl, pH 8.5. Following 6 h at room temperature, AlexaFluor-680-conjugated fibrinogen was purified from the reaction mixture by dialyzing in MWCO 30,000 Da dialysis tubing against 50 mM Tris, 150 mM NaCl, pH 7.4 for 24 h with regular replacement of buffer. 2.6. Short-term degradation under high plasmin conditions Retention of aprotinin-a2PI1e8 and WT aprotinin within fibrin matrices was examined by a two-part, short-term degradation assay at high plasmin concentration (0.1 U plasmin/100 mL fibrin gel). Fibrin matrices were made as previously specified. In the first part, matrices were directly transferred into 4 volumes of PBS with plasmin, where they were allowed to digest at 37  C for 24 h. Digestion was followed by a 24 h wash in 10 volumes of deionized water with two buffer changes to remove excess salt. After the final wash, diameters were measured for each matrix. For each condition, 6 matrices were pooled and lyophilized, followed by measurement of their collective dry weight. The experiment was repeated once more under the same conditions, but including an initial 24 h pre-wash in 4 volumes of PBS prior to plasmin degradation.

2. Materials and methods 2.7. Long term degradation under physiological plasmin conditions 2.1. Cell isolation and culture Human dermal fibroblasts (HDFs, a kind gift from the laboratory of Prof. Melody Swartz, Ecole Polytechnique Fédérale de Lausanne) were isolated from neonatal foreskin and were expanded in complete medium comprised of Ham’s F10 (Invitrogen/Gibco, Basel, CH) supplemented with 10% fetal bovine serum (Invitrogen/ Gibco, Basel, CH), 1% penicillin/streptomycin, and 0.25 mg/mL fungizone amphotericin B (Invitrogen/Gibco, Basel, CH). Cells were utilized at passage 9. Medium was changed every second day. 2.2. Recombinant aprotinin-a2PI1e8 production Complimentary DNA (cDNA) encoding for bovine aprotinin was PCR-amplified from the plasmid pRK20.1.2 (a kind gift from Bayer), using primers specific for insertion within the multiple cloning site of the expression vector pGEX-4T-1 (GE Healthcare, Chalfont St. Giles, UK). The 30 primer was designed such that the C-terminus of the amplified cDNA included the transglutaminase substrate sequence, NQEQVSPL. The ligation product, herein termed pGEX-aprotinin-a2PI1e8, provided for protein expression via the glutathione S-transferase (GST) gene fusion system. The fusion protein, herein termed GST-aprotinin-a2PI1e8 was expressed in BL21 Escherichia coli (E. coli) and purified by GST affinity chromatography (AKTAFPLC, GE Healthcare, Chalfont St. Giles, UK). Liberation of aprotinin-a2PI1e8 from the fusion system was achieved by digestion with bovine thrombin (Sigma-Aldrich, Buchs, Switzerland) followed by GST affinity chromatography. Endotoxin was removed via extensive column washing with Triton X-114. Protein identity was confirmed by tandem mass spectrometry, and purity was verified by SDS-PAGE. 2.3. Plasmin inhibition assay Bioactivity was assessed via a plasmin inhibition assay. Specified concentrations of aprotinin-a2PI1e8 or WT aprotinin were incubated with 0.1 U/mL human plasmin (Roche, Rotkreuz, CH) in the presence of 100 mM fluorogenic plasmin substrate, N-Succinyl-Ala-Phe-Lys-7-amido-4-methylcoumarin acetate salt (Sigma-Aldrich) in phosphate-buffered saline (PBS) at pH 8.5. Fluorescence emission was measured at 438 nm with a plate reader (Safire II, Tecan Sales Switzerland AG, Mannedorf, CH), relative fluorescence between control and mutant were compared. 2.4. Fibrin gel preparations Fibrin matrices were made by mixing 4 mg/mL human fibrinogen (plasminogen, von Willebrand factor-, and fibronectin-depleted; Enzyme Research Laboratories, South Bend, IN, USA), 1 U/mL factor XIIIa (gift of Baxter BioSurgery, Vienna, Austria), 2 U/mL human thrombin (Sigma-Aldrich, Buchs, Switzerland), and 2.5 mM Ca2þ in Tris-buffered saline (TBS). Matrices containing aprotinin-a2PI1e8, WT aprotinin, or no inhibitor (TBS) were obtained by adding the specified concentrations of inhibitor or TBS to the constituents, respectively. Matrices were allowed to polymerize at 37  C for 45 min before being transferred into medium.

To model in vivo plasmin conditions, fluorescently-labeled fibrin matrices containing aprotinin-a2PI1e8, WT aprotinin, or no inhibitor were exposed to physiological levels of plasmin (2.4 nM) [14] for 2 wk. Media was collected for analysis and replaced daily. Fluorescence of media was determined by measuring the emission at 516 nm. Western blots were performed by resolving proteins on 12% SDS-PAGE gels, followed by protein transfer onto membranes (ImmobilonTM transfer membranes, Milian, Geneva, CH). Membranes were blocked in a solution containing 10% milk in TBS with 0.1% tween-20 (TBST) at 4  C overnight. Membranes were washed with TBST and incubated with a goat anti-human fibrinogen antibody coupled to horseradish peroxidase (1:5000 dilution, MP Biomedicals, Basel, CH) at room temperature for 1 h. Membranes were washed and drained of excess buffer, and proteins were detected using a chemiluminescent substrate (SuperSignalÒ West Pico Chemiluminescent Substrate, Thermo Fisher Scientific, Lausanne, CH). Our laboratory-formulation fibrin matrices, hereafter referred to simply as “fibrin,” were compared with commercial fibrin sealants Omrixil (Johnson & Johnson) and Tisseel (Baxter) in a plasmin challenge at physiological plasmin levels. Tisseel and Omrixil matrices were made by mixing equal parts of their dual solutions (as specified by the manufacturers), and supplemented with inhibitor variant, inhibitor, or PBS to obtain matrices containing 2.61 mM aprotinin-a2PI1e8, 2.61 mM WT aprotinin, or no inhibitor, respectively. Matrices were placed in 10 volumes of PBS with physiological concentrations of plasmin and were maintained in 37  C incubators for the duration of the experiment. Media was collected and refreshed at each time point. Time points were taken every 8 h for the first 3 d and every 24 h thereafter until 21 d. Photographs were taken daily to monitor macroscopic matrix appearance. At the end of the 21 d, remaining matrices were pooled and lyophilized, followed by measurement of their collective dry weight. Selected time points were evaluated by Western blot, performed as previously described. 2.8. Cell proliferation and cell-induced matrix degradation Fibrin matrices were made as previously described, except for addition of cells to aforementioned constituents. HDFs were embedded within matrices prior to gelation at a concentration of 14,000 cells/mL. Cell proliferation within fibrin matrices was assessed via metabolic activity indicated by AlamarBlue, as per the manufacturer’s instructions (Serotec Ltd., Dusseldorf, DE). Medium was removed from all wells containing matrices and was replaced with fresh complete medium. AlamarBlue was mixed into each well to attain 10% by volume concentration, followed by incubation at 37  C for 2 h. Fluorescence was measured at 590 nm with a plate reader. Metabolic activity of cells is measured by an oxidation-reduction indicator that fluoresces in response to chemical reduction of medium resulting from cell growth. All samples were normalized against a negative control containing a polymerized fibrin gel, medium, and AlamarBlue only. Proliferation and diameter measurements were performed with 6 gels at each condition. After 2 wk, histology was performed by fixing matrices in 2% paraformaldehyde and cryosectioning 30 mm slices. Microscopy images were taken on a fluorescence microscope, using a 20 objective (Leica).

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2.9. Subcutaneous matrix implantation All animal procedures were previously approved by the Swiss Vaudoise Veterinary Office. Female C57BLKS/J mice, 8e10 wk old were used for this study. All materials were prepared prior to surgery. Fibrin matrix components contained 10 mg/mL Alexafluor-680-labeled fibrinogen, 1 U/mL factor XIIIa, 2 U/mL human thrombin, and 2.5 mM Ca2þ in TBS. Matrices were supplemented with inhibitor variant, inhibitor, or PBS to obtain matrices containing 25 mM aprotinin-a2PI1e8, 25 mM WT aprotinin, or no inhibitor, respectively. Components were mixed immediately prior to use. Animals were anesthetized with 4% induction of isoflurane followed by maintenance at 2%. The dorsal skin was shaved, disinfected with Betadine, followed by 70% ethanol. Two incisions, approximately 1.5 cm long, were made with a scalpel along the left and right sides of the dorsum. A cylindrical mold was placed within the spreadskin following theincision, exposing a circular pocket over the muscle tissue with a 7 mm diameter. Gels were polymerized in situ for approximately 3 min before removal of the mould and closure of the wound with 4.0 nondegradable ProleneÒ sutures (Johnson & Johnson, Spreitenbach, CH), which also served as locational markings. The animals were euthanized after 0, 5, 10, 14, 18, and 24 d by means of carbon dioxide asphyxiation. Gels were excised together with the surrounding skin and connective tissue. Of the 6 resected per condition, 2 were submitted for histology and 4 were homogenized in M-PER buffer (Thermo Scientific) (0.5 mg tissue/mL solution). Histology was performed with Masson’s trichrome staining, and immunohistochemistry was performed with goat anti-human fibrinogen antibody (1:400 dilution, MP Biomedicals, Basel, CH) and DAPI nuclear staining. All slides were imaged on a Leica microscope with a 10 objective. Homogenized tissue samples were resolved on a 12% SDS-PAGE gel, followed by scanning at 700 nm wavelength with an LI-COR Odyssey imaging system. 2.10. Statistics All experimental data represent mean  standard deviation of measurements. Student’s two-tailed t-test was used for comparison of groups against negative controls, using GraphPad Prism 4.0.

removed by column washing with Triton X-114 during affinity chromatography, before cleavage of aprotinin-a2PI1e8 from the glutathione S-transferase fusion by thrombin. The protein identity was verified with tandem mass spectrometry and its purity was assessed by SDS-PAGE (Fig. 1A). The primary concentration of aprotinin utilized in our studies was 2.61 mM. This is the equivalent to 100 kallikrein inhibitor units (KIU)/mL of WT aprotinin (Roche) and thus can be readily compared against concentrations used in clinical applications, normally reported in KIU/mL. For example, CSL Behring’s Beriplast and Baxter’s Tisseel fibrin sealants contain 500 and 1500 KIU/mL aprotinin, respectively, according to product package inserts. Furthermore, this concentration has been previously used by our laboratory in in vitro cell culture [15] and compares well with concentrations successfully used by others in similar in vitro applications [8,16,17]. In order to determine the bioactivity of the bacteriallyexpressed aprotinin variant, inhibitory function was assessed via a plasmin substrate cleavage assay, whereby aprotinin-a2PI1e8 was found to be as active as WT aprotinin at tested concentrations (Fig. 1B). Care was taken to add plasmin simultaneously to all reactions and to minimize time between plasmin addition and fluorescence readings. Plasmin was added to all reaction mixtures and resulted in cleavage of the fluorogenic plasmin substrate only in reactions lacking plasmin inhibitors. Reactions containing either aprotinin-a2PI1e8 or WT aprotinin did not result in substantial cleavage of plasmin substrate.

3. Results

3.2. Functional retention of aprotinin-a2PI1e8 within fibrin

3.1. Generation of aprotinin variant, aprotinin-a2PI1e8

Retention of aprotinin-a2PI1e8 within the fibrin matrix was demonstrated using a functional fibrinolysis assay. Fibrin matrices containing either 2.61 mM of aprotinin-a2PI1e8, WT aprotinin, or no inhibitor were immediately exposed to a high level of plasmin (0.1 U plasmin/100 mL gel) for 24 h to test inhibitor effectiveness against plasmin degradation. Gels were extensively washed in deionized water to remove remaining plasmin, degraded fibrin, and buffer salts before being pooled, lyophilized, and weighed. Dry weight of lyophilized gels revealed that fibrin matrices containing aprotinin-a2PI1e8 retained 80% of their mass and those containing WT aprotinin retained only 50%. As expected, fibrin matrices containing no inhibitor completely degraded during the 24 h exposure to plasmin (Fig. 2A).

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We applied a protein modification scheme previously developed in our laboratory [13] to generate an aprotinin variant, aprotinin-a2PI1e8, that would allow for its covalent incorporation into fibrin matrices during normal thrombin and factor XIIIamediated polymerization. Covalent incorporation was mediated by fusion of a factor XIIIa substrate sequence tag, derived from the N-terminal 8 residues of a2-plasmin inhibitor, to the C-terminus of aprotinin. Using standard cloning techniques, aprotinin cDNA was inserted into an expression vector with addition of the substrate sequence tag. Aprotinin-a2PI1e8 was then produced in E. coli as a fusion protein with glutathione S-transferase. Endotoxin was

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Fig. 1. Purification and characterization of recombinant aprotinin-a2PI1e8. GST-fused aprotinin-a2PI1e8 was purified via glutathione affinity chromatography, enzymatically digested to remove the GST tag, and re-purified on a glutathione column to obtain a pure product. A) Coomasie-stained SDS-PAGE analysis of the purified recombinant protein shows a slight increase in molecular weight as compared to naturally derived WT aprotinin, due to the transglutaminase substrate sequence fusion. Bioactivity of the aprotinin variants was assessed by measuring their ability to inhibit the cleavage of the model plasmin substrate N-succinyl-Ala-Phe-Lys-7-amido-4-methylcoumarin in the presence of 0.1 U/mL plasmin, and either 2.61 mM (B) or 261 nM (C) WT aprotinin or aprotinin variant. WT and aprotinin-a2PI1e8 exhibited comparable inhibitory characteristics, confirming that the bacteriallyexpressed aprotinin with an 8 amino acid C-terminal factor XIIIa substrate tag has comparable biochemical characteristics as the naturally derived WT aprotinin. B) and C) values are reported as mean  S.D., N ¼ 2.

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wash, whereas the fibrin matrices containing WT aprotinin decreased in diameter by 13% without the pre-wash.

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Fig. 2. Fibrin matrix integrity following in vitro plasmin challenge. The structural integrity of fibrin gels containing either no inhibitor, WT aprotinin, or aprotinin-a2PI1e8 following 24 h incubation in 0.1 U/mL plasmin was assessed by measuring matrix mass or diameter. A) Directly following gel formation, matrices were placed in plasmincontaining buffer for 24 h at 37  C (open bars). Aprotinin-a2PI1e8-protected the matrices to a greater extent than WT aprotinin, evidenced by the maintenance of matrix mass. To assess whether the inhibitor was bound within the matrix, matrices were washed for 24 h in 10 volumes of PBS, then transferred to plasmin-containing buffer for 24 h at 37  C (filled bars). The protective effect of WT aprotinin as previously seen (open bars) was completely abolished, whereas aprotinin-a2PI1e8-containing matrices were still present at high structural integrity. B) Measurement of matrix diameter under the same conditions as described above corroborated the results seen by measurement of mass. The results also suggest that plasmin-mediated proteolysis occurred uniformly throughout the bulk of the matrix.

To determine the effects of inhibitor retention or diffusive release on fibrin protection, the same steps were repeated; however, a 24 h pre-wash step was included to allow diffusive release of unbound inhibitor before exposure of the gels to plasmin. After the pre-wash, fibrin matrices containing WT aprotinin or no inhibitor degraded completely during subsequent exposure to plasmin for 24 h, whereas fibrin matrices containing aprotinin-a2PI1e8 retained 63% of their mass (Fig. 2B). In both assays, diameter measurements corroborated the trend in mass measurements, although differences between groups were reduced (Fig. 2C). Fibrin matrices containing aprotinin-a2PI1e8 decreased in diameter by 4% without the pre-wash and by 17% with the pre-

To model in vivo plasmin conditions, fluorescently-labeled fibrin matrices containing aprotinin-a2PI1e8, WT aprotinin, or no inhibitor were exposed to physiological amounts of plasmin [14] for 14 d. Media was collected for analysis and replaced daily. Fluorescence of fibrin degradation products within the media was quantifiable, and degradation exhibited similar yet delayed kinetics to the previous 24 h fibrinolysis study. Fibrin matrices containing no inhibitor degraded completely by day 5, whereas those containing 2.61 mM and 25 mM WT aprotinin degraded by day 6 and day 9, respectively (Fig. 3A). However, fibrin matrices containing aprotinin-a2PI1e8 degraded only minimally after 14 d at both of the investigated concentrations. Anti-fibrinogen Western blots of media taken from select time points confirmed the plasmin degradation trend, demonstrated at early stages by prominent bands at 43 kD, and later by additional bands at 58 kD and w100 kD (Fig. 3B). Molecular weights of plasmin degradation products were in keeping with previously reported sizes [18,19]. Commercial fibrin sealants Omrixil (Johnson & Johnson) and Tisseel (Baxter), as well as fibrin gels made from plasminogen (and other factor)-depleted fibrinogen, were combined with supplements of aprotinin-a2PI1e8, WT aprotinin, or PBS (no inhibitor). For simplicity, these matrices are referred to as Omrixil, Tisseel, and fibrin matrices, respectively, regardless of the universal presence of fibrin within all investigated matrices. In all cases, matrices containing WT aprotinin or no inhibitor showed extreme degradation by day 5, as demonstrated by macroscopic analysis and supporting anti-fibrinogen Western blot (Fig. 4A), and degraded completely by day 6. Initial addition of WT aprotinin made little difference in matrix degradation rates, as compared to matrices containing no inhibitor. In contrast, all matrices with addition of aprotinin-a2PI1e8 sustained exposure to physiological levels of plasmin for longer periods of time, ranging from approximately twice as long as no inhibitor controls, for Omrixil and Tisseel matrices, to more than three times as long as no inhibitor controls for fibrin matrices (Fig. 4B). By 21 d, fibrin matrices containing aprotinin-a2PI1e8 were the only matrices which had not completely degraded, and moreover, these matrices showed few signs of degradation, retaining 78% of their original mass. 3.4. Matrix longevity extension in cell culture The capacity of fibrin matrices to support cell growth was assessed by characterization of fibrin matrix diameter, cell proliferation as measured by cellular metabolism, and histology. Fibrin matrices containing aprotinin-a2PI1e8, WT aprotinin, or no inhibitor were seeded with HDFs and cell proliferation was monitored over the course of 28 d. Diameters of matrices were measured once a week so as to minimize gel handling, and histology was performed at day 14. Fibrin matrices containing WT aprotinin and no inhibitor displayed similar characteristics; matrices were contracted by cells over time and eventually degraded by day 21 under both conditions (Fig. 5A). Fibrin matrices containing aprotinin-a2PI1e8, however, were contracted by cells more slowly and maintained the capacity to support cell growth for at least 28 d. Furthermore, in fibrin matrices containing WT aprotinin or no inhibitor, cellular metabolism peaked at day 10 and steadily declined thereafter, whereas fibrin matrices containing aprotinin-a2PI1e8 continued to support increasing cell proliferation for 18 d and exhibited only a slow decline in cell activity thereafter (Fig. 5A). Histology sections after 14 d revealed numerous

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Day Fig. 3. Assessment of fibrinolysis in aprotinin-containing matrices. Fibrinogen was covalently modified with FITC and purified prior to matrix formation. Following matrix formation in the presence of varying concentrations of either WT or aprotinin-a2PI1e8 or no inhibitor, matrices were incubated in a physiological concentration (2.4 nM) of human plasmin for 14 d at 37  C. Every 24 h, the matrix-containing buffer was analyzed for presence of soluble, and therefore cleaved, fibrin by measuring FITC fluorescence at 516 nm (A) and by Western blot (B), and the matrices were replenished with fresh plasmin-containing buffer. A) Matrices containing either no inhibitor or WT aprotinin released much greater amounts of soluble fibrin over 14 d of plasmin challenge as compared to aprotinin-a2PI1e8-containing matrices. B) Anti-fibrinogen Western blot analysis corroborated the fluorescence measurements in that no staining, and thus no detectable cleaved fibrin, was present in samples taken from buffer of matrices containing aprotinin-a2PI1e8. Notably, even at a 10-fold higher concentration of 2.61 mM, WT aprotinin was unable to offer protection from plasmin comparable to aprotinin-a2PI1e8 at a 10-fold lower concentration of 261 nM, highlighting the importance of covalent attachment in maintaining inhibitory potency. A) Values are reported as mean  S.D., N ¼ 6.

perforations in fibrin matrices containing WT aprotinin or no inhibitor, whereas cavities were not detectable in fibrin matrices containing aprotinin-a2PI1e8 (Fig. 5C). Microscopic evidence of HDFs along perforation perimeters suggests perforations were created by induced local cell degradation. 3.5. Fibrin implant longevity extension in vivo The ability of inhibitors to prolong the presence of fibrin matrices in vivo was investigated by subcutaneously implanting fluorescently-labeled fibrin matrices with aprotinin-a2PI1e8, WT aprotinin, or no inhibitor under the back skin of C57/BL6J mice. Six samples for each inhibitor were harvested at days 0, 5, 10, and 14. From each set of 6 samples, 2 were used for histology (Fig. 6A) and 4 were homogenized and analyzed by infrared scanned SDS-PAGE (Fig. 6B). All fibrin matrices were readily detectable at day 5 by both histological and infrared SDS-PAGE visualization. By day 10, fibrin matrices containing WT aprotinin or no inhibitor were no longer detectable by either method, though fibrin matrices containing aprotinin-a2PI1e8 were clearly visible via histology and infrared SDS-PAGE visualization at days 10 and 14. Subsequently, the presence of fibrin matrices containing aprotinin-a2PI1e8 was explored at two additional time points: day 18 and day 24. At these late time points, fibrin matrices were detectable in all samples via the described methods; however, 2 of the 6 matrices at day 24 were not macroscopically visible at day 24 and were only faintly detectable by infrared scanned SDS-PAGE. This study demonstrated that fibrin gels containing aprotinin-a2PI1e8 can be detected in vivo for 2e4 times longer than fibrin gels containing WT aprotinin or no inhibitor. 4. Discussion The goal of this study was to develop a protease inhibitor variant that would confer extended longevity to fibrin matrices, especially in vivo. Towards this end, we designed a recombinant inhibitor mutant that would be covalently immobilized within fibrin matrices during thrombin and factor XIIIa-mediated polymerization, and thereby would provide a sustained local inhibitor reservoir near degradable fibrin sites. Aprotinin was chosen as the core inhibitor of our design due to its well-characterized behavior, simple nonglycosylated structure, and capacity to be produced in functional form in E. coli at high yields. Thus, our fibrin matrices engineered for in vivo longevity contain a naturally inspired mutant

plasmin inhibitor immobilized to the matrix during a precisely defined in situ gelation reaction. Standard cloning techniques were used to fuse a transglutaminase substrate sequence tag to aprotinin and thereby create a protease inhibitor variant, aprotinin-a2PI1e8. Because the substrate sequence tag was derived from a2-plasmin inhibitor, aprotinin-a2PI1e8 would be crosslinked to the alpha chain of fibrin, the naturally evolved site of the native inhibitor [20]. This form of biomimicry would theoretically be advantageous because aprotinin-a2PI1e8 would consequently be presented in proximity of plasmin sensitive sites on the alpha chain, of which twelve exist [21]. A bacterial expression system was successfully employed to produce an active form of aprotinin-a2PI1e8 (Fig. 1). To simplify production, we chose to bypass protein refolding steps by isolating aprotinin-a2PI1e8 only from soluble bacterial lysate and were able to extract approximately 0.6 mg fusion protein/L of culture. Additional extraction from bacterial inclusion bodies would result in substantial increase in protein yield, and previous studies have shown that complete activity recovery of reduced aprotinin is possible through subsequent refolding steps [22,23]. Aprotinin-a2PI1e8 exhibited similar activity to the WT aprotinin at the tested plasmin concentrations (Fig. 1B and C). Because plasmin is a broad-specificity protease, it is not surprising that plasmin normally circulates in blood as plasmin/antiplasmin complexes at an extremely low baseline concentration of 2.4 nM in humans [14]. However, plasminogen, the zymogen of plasmin, circulates in plasma at an almost 1000 fold higher concentration, w1.5 mM, and thus can lend to substantial, rapid increases in plasmin concentration at wound sites [24]. Our bioactivity assay demonstrated the capacity of aprotinin-a2PI1e8 and WT aprotinin to inhibit plasmin at the maximum physiologically relevant concentration of 1.5 mM. Retention of bound but not free protease inhibitor was functionally demonstrated by washing fibrin matrices supplemented with aprotinin variants for 24 h before exposure to high concentrations of plasmin. Although the pre-wash completely abolished the ability of WT aprotinin to protect fibrin matrices, aprotinina2PI1e8-containing matrices retained most of their structural integrity (Fig. 2A and B). These data suggest that immobilization of inhibitor within fibrin matrices is especially relevant for protection against proteolytic degradation. In contrast, although fibrin matrices quickly lost mass, they retained the majority of their diameter until complete degradation (Fig. 2C). This was the expected trend in fibrin matrix degradation, as plasmin cleavage

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Fig. 4. Stability assessment of commercial fibrin products. Matrices of the commercially available fibrin-based clinical sealants Omrixil and Tisseel were formed as per the manufacturer’s instructions, with the addition of either WT or aprotinin-a2PI1e8 to the formulation prior to gelation. Assessment of aprotinin-a2PI1e8’s capacity to offer enhanced protection from plasmin to clinically-used fibrin products was conducted by a time-course plasmin challenge (2.4 nM plasmin, 2.61 mM WT aprotinin or aprotinin variant). Our laboratory-formulation of a fibrin gel was used as a reference. (A) Photographs taken of the gels before complete degradation show that aprotinin-a2PI1e8 effectively increases the lifespan of the fibrin product over two-fold. Displayed time points were the latest days on which remaining matrices for all aprotinin conditions provided sufficient contrast for image visibility. Images above each photo represent the 43 kDa major cleavage component band observed following anti-fibrinogen Western blot analysis of the matrix buffer component. Well diameter ¼ 97 mm. (B) Under the conditions described above, the maximum time point (in days) at which residual fibrin could be observed.

has previously been described in detail to occur in bulk degradation throughout the matrix rather than as surface erosion [25]. When exposed to physiological concentrations of plasmin for 14 d, aprotinin-a2PI1e8 again demonstrated the capacity to protect fibrin matrices from degradation better than WT aprotinin (Fig. 3A and B). Notably, at the lower concentration of 2.61 mM, WT aprotinin prolonged the presence of the fibrin matrices for only one

additional day, and a further almost 10-fold increase in the concentration of WT aprotinin from 2.61 mM to 25 mM prolonged fibrin matrix presence for a maximum of only 3 d relative to controls with no inhibitor. In contrast, fibrin matrices containing aprotinin-a2PI1e8 showed minimal signs of degradation after 2 wk at all concentrations tested. Thus, these data further highlighted the importance of covalent inhibitor attachment in prolonging the

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Fig. 5. Effect of aprotinin variants on 3D culture of human dermal fibroblasts in fibrin matrices. When seeded with HDFs, fibrin matrices containing aprotinin-a2PI1e8 retained their structure to a higher degree than matrices containing either no inhibitor or WT aprotinin. A) Cell-mediated gel degradation was measured by monitoring gel diameter over 28 d in culture. Fibrin aprotinin-a2PI1e8-supplemented matrices retained significantly larger diameters than WT aprotinin-supplemented matrices (P < 0.0001) B) Cellular proliferation was measured over the same time period, which demonstrated that aprotinin-a2PI1e8-supplemented matrices were able to sustain and conduct HDF growth over markedly longer durations than WT aprotinin-containing matrices. Visual confirmation of aprotinin-a2PI1e8 protection from cell-mediated proteolysis was confirmed microscopically following 14 d of culture. HDFs cultured in matrices containing no inhibitor (C) or WT aprotinin (D) degraded large areas of matrix, evidenced by the large holes visible by phase contrast microscopy. In contrast, aprotinin-a2PI1e8 matrices (E) retained uniform structural integrity while still conducting the growth of healthy HDFs, as evidenced by their elongated morphology. C), D), and E) scale bar ¼ 200 mm. A) and B) Values are reported as mean  S.D., N ¼ 6.

durability of fibrin matrix and also served as a promising indicator of possible therapeutic potential in vivo. Supplementation of aprotinin-a2PI1e8 into commercial fibrin sealants Omrixil and Tisseel more than doubled their durability when compared to WT aprotinin, thus reconfirming the ability of aprotinin-a2PI1e8 to confer extended longevity to fibrin matrices (Fig. 4a and b). Additionally, these results demonstrate the immediate therapeutic potential of aprotinin-a2PI1e8 as a simple additive to current, clinically implemented fibrin sealant products. As expected, Omrixil matrices, which do not contain protease inhibitors within their normal formulation, degraded more quickly than Tisseel matrices, which contain a mean aprotinin concentration of 1500 U/mL. However, of the aprotinin-a2PI1e8-containing matrices, we found it surprising that Omrixil and Tisseel matrices degraded significantly more quickly than our laboratory formulated fibrin matrices, which contain approximately ten times less fibrinogen, yet retained the bulk of their mass (78%) at 21 d. One explanation for this discrepancy may stem from suboptimal mixing of the two highly viscous components, which are combined to form each commercial sealant. Notably, insufficient mixing of the constituents in opposing component solutions, such as fibrinogen and thrombin, could result in heterogeneous fibrin polymerization and perhaps weaken the overall matrix structure. Similarly, insufficient mixing of opposing constituents, factor XIII and its required cofactor, calcium, might also lend to suboptimal crosslinking of the fibrin matrices [26]. Another explanation for the discrepancy may be the

inclusion of plasminogen, the zymogen of plasmin, within the commercial fibrin sealants’ components. For example, the Tisseel product specification states that the final concentration of plasminogen within the sealant may be as high as 0.65 mM and that other unspecified proteins contribute additionally to the clot. Small amounts of tissue plasminogen activator (t-PA) and urokinase plasminogen activator (u-PA), which may be present within the unspecified proteins, could collectively contribute to a relevant enhancement in fibrin degradation [25]. More specifically, t-PA and plasminogen bind to the same set of sites in fibrin during polymerization, and upon binding, undergo conformational changes which result in hundred-fold enhancement of plasminogen activation, and a subsequent positive feedback loop markedly accelerates cleavage of fibrin. When tested in cell culture conditions, aprotinin-a2PI1e8-containing fibrin matrices supported HDF growth at least 33% longer than matrices containing WT aprotinin or no inhibitor (Fig. 5A and B). Interestingly, the presence of aprotinin-a2PI1e8 seemed to prevent the extensive cell-localized matrix degradation which was apparent after 2 wk in fibrin matrices containing WT aprotinin or no inhibitor (Fig. 5C). This phenomenon may have been due to inhibition of plasmin as well as inhibition of pro-MMPs in the vicinity of cells by the immobilized aprotinin-a2PI1e8 still present at later time points. More specifically, plasmin has been reported to either directly or indirectly activate certain pro-MMPs, such as MMP-1, -2, -3, and -9, [27e29] and these MMPs have been shown,

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Fig. 6. Stability of aprotinin-a2PI1e8-supplemented matrices in vivo. A) Histological analysis of subcutaneously implanted fibrin matrices containing no inhibitor, WT aprotinin, or aprotinin-a2PI1e8 revealed an extensive increase in gel longevity for matrices containing aprotinin-a2PI1e8. Immunohistochemical staining with anti-fibrinogen antibody both confirmed the presence of the original implanted fibrin and indicated its structural boundaries. Following 10 d in mice, only aprotinin-a2PI1e8-protected matrices were visible, and remained present for at least 24 d. B) AlexaFluor-680-conjugated fibrin matrices were implanted subcutaneously in mice and resected for analysis at varying time points. Following digestion and homogenization, the gel sample was analyzed by SDS-PAGE and imaged on a fluorescence gel scanner. Each lane represents one implant. Similar to histological and immufluorescence results, at d 10 no fluorescent signal from remaining fibrin matrix was visible except in matrices containing aprotinin-a2PI1e8. Aprotinin-a2PI1e8-protected matrices remained visible by fluorescence following 24 d post implantation. Longer time points were not evaluated. A) scale bar ¼ 1 mm.

in turn, to be secreted by HDFs [30,31] and linked to fibrin degradation [8,28,32]. Consequently, partially blocking these activities may have resulted in fibrin matrices containing aprotinin-a2PI1e8 to maintain structural integrity while still permitting embedded HDFs to spread, as evidenced by histology after 14 d, and to remain metabolically active for at least 28 d, as evidenced by AlamarBlue. Aprotinin-a2PI1e8 was also able to extend the durability of fibrin matrices by more than two-fold compared to WT aprotinin in vivo as demonstrated by subcutaneous implantation of fibrin matrices

into mice (Fig. 6A and B). Although fibrin matrices containing WT aprotinin or no inhibitor were no longer detectable by day 10, aprotinin-a2PI1e8-protected fibrin matrices were still detectable in vivo at 24 d. Histology of sectioned implant areas revealed minimal inflammatory cell infiltration into fibrin matrices. This could be due to the decrease in fibrin degradation and concomitant decrease in locally released fibrin degradation products, which normally serve as mitogenic factors for a variety of cell types, including endothelial cells, fibroblasts, smooth muscle cells, and lymphocytes [33,34].

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Low inflammatory cell presence could also be explained by the lack of cell signaling molecules, such as growth factors and cytokines, which would be released as inflammatory cues from physiological blood clots but were not present at relevant levels in the factordepleted fibrin matrices. Consequently, fibrin matrices containing aprotinin-a2PI1e8 were both long-lasting and well-tolerated as subcutaneous implants in vivo. 5. Conclusion We conclude that our recombinant protease inhibitor variant, aprotinin-a2PI1e8, can confer extended longevity to fibrin matrices more effectively than WT aprotinin in vitro and in vivo. Our protein modification scheme generated a small, nonglycosylated inhibitor variant that can be covalently immobilized within fibrin matrices during normal thrombin and factor XIIIa-mediated polymerization. A plasmin inhibition assay indicated that aprotinin-a2PI1e8 and WT aprotinin exhibited similar bioactivity. However, aprotinin-a2PI1e8containing matrices greatly out performed WT aprotinin-containing matrices when challenged with physiological plasmin levels for 2 wk, thus highlighting the importance of inhibitor immobilization for protection against matrix degradation. Plasmin challenge of commercial fibrin sealants Omrixil and Tisseel, which were supplemented with aprotinin-a2PI1e8, WT aprotinin, or no aprotinin, further corroborated these results. Moreover, by comparison of aprotinin-a2PI1e8 with wild type (WT) aprotinin in in vitro and in vivo settings, we demonstrated that our aprotinin variant is more effective than the WT form of the protein at extending longevity of fibrin matrices. Premature fibrinolysis and elevated protease inhibitor doses are two clinically important shortcomings of many fibrin matrices used in surgery. An inhibitor that begins to surmount these shortcomings by extending fibrin matrix longevity while reducing inhibitor dosage may be a useful tool for extending the applicability of fibrin-based biomaterials in surgery and tissue engineering. Acknowledgments The authors would like to acknowledge the EPFL Proteomics Core Facility for tandem mass spectrometry, the EPFL Histology Core Facility for histological sectioning and staining, the EPFL BioImaging and Optics Platform facility for microscopy support, Bayer for aprotinin cDNA, Baxter Biosurgery for factor XIIIa, and EuroSTEC (Soft Tissue Engineering of Congenital Birth Defects in Children; E.U. FP6: reference LSHB-CT-2006-037409) for partial funding. Appendix Figures with essential color discrimination. Figs. 4 and 6 in this article are difficult to interpret in black and white. The full color images can be found in the on-line version, at doi:10.1016/j. biomaterials.2010.08.109. References [1] Spotnitz WD. Fibrin sealant: past, present, and future: a brief review. World J Surg 2010;34:632e4. [2] Fu JZ, Li J, Yu ZL. Effect of implanting fibrin sealant with ropivacaine on pain after laparoscopic cholecystectomy. World J Gastroenterol 2009; 15:5851e4. [3] Danielsen PL, Agren MS, Jorgensen LN. Platelet-rich fibrin versus albumin in surgical wound repair: a randomized trial with paired design. Ann Surg 2010;251:825e31. [4] Garcia-Olmo D, Herreros D, Pascual I, Pascual JA, Del-Valle E, Zorrilla J, et al. Expanded adipose-derived stem cells for the treatment of complex perianal fistula: a phase II clinical trial. Dis Colon Rectum 2009;52:79e86. [5] Ahmed TA, Dare EV, Hincke M. Fibrin: a versatile scaffold for tissue engineering applications. Tissue Eng Part B Rev 2008;14:199e215.

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