Enzymatically-responsive pro-angiogenic peptide-releasing poly(ethylene glycol) hydrogels promote vascularization in vivo

Abstract

Therapeutic angiogenesis holds great potential for a myriad of tissue engineering and regenerative medicine approaches. While a number of peptides have been identified with pro-angiogenic behaviors, therapeutic efficacy is limited by poor tissue localization and persistence. Therefore, poly(ethylene glycol) hydrogels providing sustained, enzymatically-responsive peptide release were exploited for peptide delivery. Two pro-angiogenic peptide drugs, SPARC₁₁₃ and SPARC₁₁₈, from the Secreted Protein Acidic and Rich in Cysteine, were incorporated into hydrogels as crosslinking peptides flanked by matrix metalloproteinase (MMP) degradable substrates. In vitro testing confirmed peptide drug bioactivity requires sustained delivery. Furthermore, peptides retain bioactivity with residual MMP substrates present after hydrogel release. Incorporation into hydrogels achieved enzymatically-responsive bulk degradation, with peptide release in close agreement with hydrogel mass loss and released peptides retaining bioactivity. Interestingly, SPARC₁₁₃ and SPARC₁₁₈-releasing hydrogels had significantly different degradation time constants in vitro (1.16 and 8.77 x10⁻² hour⁻¹, respectively), despite identical MMP degradable substrates. However, upon subcutaneous implantation, both SPARC₁₁₃ and SPARC₁₁₈ hydrogels exhibited similar degradation constants of ~ 1.45 x10⁻² hour⁻¹, and resulted in significant ~ 1.65- fold increases in angiogenesis in vivo compared to controls. Thus, these hydrogels represent a promising pro-angiogenic approach for applications such as tissue engineering and ischemic tissue disorders.

Graphical Abstract

1. Introduction

Therapeutic angiogenesis has great promise for regenerative medicine applications, including treatment of ischemic tissue disorders [1] and engineering tissues that are > 100–200 μm thick [2]. Proangiogenic approaches have been developed, often utilizing delivery of angiogenic proteins [3, 4] or peptide drugs [5, 6]. While some peptides consist of entirely novel sequences [7], they typically mimic the active region(s) of larger proteins or growth factors [5, 6]. Due to low molecular weights compared to proteins (typically < 10 kDa), peptides can be produced synthetically and delivered at higher concentrations compared to protein counterparts. Additionally, peptides do not require complex tertiary structures for bioactivity, resulting in increased stability in vivo [5]. However, both proteins and peptides suffer from poor targeting, short circulation time, and rapid clearance when delivered by direct injection [8, 9], motivating the development of controlled release systems.

Current peptide delivery systems are limited [10]. Repeat injections suffer from poor patient compliance [9], osmotic pumps require revision surgeries [11], and diffusional release of therapeutics from polymers [12, 13] or hydrogels [14] result in pre-dictated release rather than delivery in response to the specific biological requirements of the tissue. To overcome hurdles associated with peptide delivery, we recently developed a poly(ethylene glycol) (PEG) hydrogel platform technology that provides enzymatically-responsive release of therapeutic peptides (Figure 1) [15]. Enzymatically-responsive hydrogel degradation and peptide release was achieved via step-growth reactions of norbornene-functionalized PEG with thiol-functionalized crosslinkers comprised of a peptide drug flanked by enzymatically-responsive linkages (C-IPES↓LRAG-peptide drug-IPES↓LRAG-C-G; degradable linker, “DL”). Previous work identified peptide drug properties that control drug delivery from these hydrogels [15].

Figure 1.

Poly(ethylene glycol) (PEG) hydrogels provide sustained, enzymatically-responsive release of pro-angiogenic peptides. PEG hydrogels crosslinked via thiol-ene photopolymerization between norbornene groups on PEG macromers and thiol-containing cysteine amino acids degrade in the presence of matrix metalloproteinases, releasing pro-angiogenic peptides in an enzymatically-responsive manner. ↓ indicates cleavage site. Not to scale.

Building upon that work, here we establish the therapeutic potential of this platform technology, providing sustained, localized pro-angiogenic peptide release via host tissue MMP activity. Specifically, delivery of SPARC₁₁₃ and SPARC_118, two pro-angiogenic peptide drugs was studied in vitro and in vivo. SPARC₁₁₃ and SPARC₁₁₈ are fragments of the Secreted Protein Acidic and Rich in Cysteine (SPARC) which is expressed during development and in wound repair [16–18]. SPARC is cleaved by a number of proteases in vivo, which releases domains with a variety of biological effects [19]. SPARC₁₁₃ and SPARC₁₁₈ are from released from the follistatin-like domain and contain the tripeptide GHK, which promotes angiogenesis in the rabbit cornea assay and accelerates dermal wound healing in mouse and rat models [18], In vitro, peptide efficacy was assessed with respect to endothelial cell tube formation. Continuous/sustained availability, mimicking controlled release, was compared to bolus delivery. Additionally, peptide dose-response bioactivities with and without residual MMP-substrates were investigated. Hydrogels were then formed and enzymatically-responsive hydrogel degradation, peptide release behavior, and in vitro efficacy were characterized. Hydrogels were fluorescently labeled and implanted subcutaneously. Live animal fluorescent imaging (IVIS) was used to track degradation and angiogenesis was assessed quantitatively via hemoglobin (Hb) measurements and qualitatively using multi-photon fluorescent microscopy.

2. Materials and methods

All materials were obtained from Sigma-Aldrich, and cells and cell culture materials were obtained from Lonza, unless otherwise noted.

2.1. Materials synthesis and characterization

2.1.1. Peptide synthesis

Peptide drugs (SPARC_113, SPARC_118, and the scrambled control) were synthesized in “native” (“N”) forms, containing only peptide drug, as well in “two-tailed” (“2T”) and “degradable linker” (“DL”) forms. The “DL” form has the peptide drug flanked on either side by the full MMP degradable linker IPES↓LRAG, and the “2T” form mimics release of drugs from hydrogels after substrate cleavage (Table 1).

Table 1.

Peptide sequences utilized. Standard amino acid abbreviations are used. ↓ indicates MMP cleavage site.

Peptide Sequence	Origin/Notes	Peptide Nickname
TLEGTKKGHKLHLDY	Basic region of SPARC [16–18]	SPARC113
KKGHK	Cu2+ binding region of SPARC [16–18]	SPARC118
GLKEQSPRKHRLG	Scrambled control peptide [15]	Scrambled
peptide drug-G	No additional Gly added to C-termini of Scrambled	“N” form
LRAG-peptide drug-IPES-G	No additional Gly added to N-termini of Scrambled	“2T” form
C-IPES↓LRAG-peptide drug-IPES↓LRAG-C-G	No additional Gly added to N-termini of Scrambled	“DL” form
C-KGKGKGKGK-C-G	Non-degradable crosslinker	NDL
CG-RGDS-G	Cell adhesion peptide	RGD
Texas Red-GGEGGEGC-G	Fluorescently labeled peptide	FLP*

All peptides were synthesized using a Liberty1 automated peptide synthesizer (CEM) on Fmoc-Gly-Wang resin (EMD), as previously described [15].

2.1.2. Peptide characterization and purification

After purification by dialysis (100–500 or 1000 MWCO depending upon peptide molecular weight; Spectrum Laboratories), peptide synthesis was confirmed using Matrix Assisted Laser Desorption Ionization-Time of Flight (MALDI-ToF) mass spectrometry (Bruker). Percent peptide was determined by measuring absorbance in ddH₂O at 205 nm (Evolution 300 UV/Vis detector, Thermo Scientific) [20], and purity (~ 90%) assessed by High Performance Liquid Chromatography (HPLC, Shimadzu Prominence) [15]. Peptides were stored at −20 °C.

2.1.3. Fluorescent labeling of peptides

To track hydrogel degradation in vivo, a fluorescently labeled peptide (FLP*) was synthesized. Unlabeled FLP (Table 1) was synthesized and cleaved from resin as described in section 2.1.1. Texas Red succinimidyl ester (Life Technologies) in HPLC grade DMF (10 mg/mL; Alfa Aesar) was added drop-wise to 10 molar excess FLP in HPLC grade DMF (10 mg/mL) containing 1 drop diisopropylethylamine. The mixture was stirred overnight at room temperature in the dark. FLP* was purified by dialysis (1000 MWCO tubing), collected by lyophilization, labeling verified by MALDI-ToF, and stored at −80 °C until use.

2.1.4. Synthesis of norbornene-functionalized poly(ethylene glycol)

Norbornene-functionalized poly(ethylene glycol) (PEGN) was synthesized from 4-arm 10 kDa PEG (JenKem Technologies USA) as previously described [21]. Functionalization was determined (> 95%) with ¹H-NMR (Bruker Avance 400) in CDCl₃, comparing peaks at δ = 6.25 – 5.8 ppm (8 H/molecule, norbornene vinyl protons, multiplet) and 4.35 – 4.05 ppm (8 H/molecule, -COOCH₂-, doublet), to those at 3.9 – 3.35 ppm (892 H/molecule, -CH₂CH₂O-, multiplet). The final product was dialyzed overnight in ddH₂O using 1000 MWCO dialysis tubing, collected by lyophilization, and stored at −20 °C.

2.2. Enzymatically-responsive hydrogel formation and characterization

2.2.1. Hydrogel formation

Enzymatically-responsive PEG hydrogels were formed as previously described (10 wt% PEGN, 1:1 thiol:ene ratio, 0.05 wt% lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) photoinitiator) [15]. For in vitro degradation and diffusive release studies, precursor solutions were injected in custom cylindrical molds, producing gels with a diameter ~ 5 mm and height ~ 2 mm. For “2T” peptide diffusive release studies, “2T” peptide was added to the precursor solution at equal molar amounts as the crosslinking peptide “NDL”. For in vivo studies, gels were polymerized within non-degradable cylindrical reactors made from pharmaceutical grade silicone tubing (Thermo Scientific) to aid in hydrogel localization (outer diameter = 7.94 mm (5/16”), inner diameter = 4.76 mm (3/16”), and height = 2.25 mm). Hydrogels for in vivo studies contained 2.8 mM RGD to facilitate cell adhesion [22] and 10 μM FLP* to facilitate non-invasive hydrogel tracking [23] in PBS. All gels were 40 μL in volume, and all precursor solutions exposed to 365 nm UV light (intensity ~ 2.5 mW/cm²) for 10 minutes for crosslinking.

2.2.2. Hydrogel degradation

After formation, hydrogels were incubated in 1 mL buffer (10 mM CaCl₂, 50 mM NaCl, 50 μM ZnCl₂ (Alfa Aesar), 50 mM Tricine (Acros Organics), and 0.05 wt% Brij35 (Alfa Aesar) in ddH₂O, pH 7.4) at 37 °C. After 24 hours, buffer solutions were exchanged with either fresh buffer or buffer containing 10 nM recombinant human MMP2 (MMP2, PeproTech). As MMP2 inactivates over time, solutions were collected and replaced at a minimum every 48 hours [24]. At various time points, solutions were collected and stored at −80 °C until quantification, and hydrogel wet and dry (post-lyophilization) masses were obtained (swelling ratio Q=M_wet/M_dry). Hydrogel degradation kinetics were fit to a pseudo-first-order degradation equation [25] to determine the degradation kinetic time constant, k_deg (eqn. 1):

$$\frac{M_t}{M_0}\propto e^{-k_{\mathit{deg}}\ast t}$$ (eqn. 1)

2.2.3. Quantification of peptide released from hydrogels

The release of tethered and encapsulated “2T” peptides was quantified by HPLC as previously described [15]. Peptide concentration was determined by comparing peak area to a standard curve formed using the “2T” version of the peptide. Encapsulated “2T” peptide diffusive release data was fit to a Fickian diffusion model and the diffusion coefficient for each gel calculated [26].

2.3. Assessment of peptide drug bioactivity in vitro

HUVECs, originally obtained from Lonza, were maintained in Endothelial Growth Media 2 (Endothelial Basal Media-2 (EBM-2) containing EGM-2 SingleQuots; EGM-2), at 37 °C with 5% CO₂. Cells were grown for at least two passages after thawing, and used prior to passage 10. Control media for all in vitro angiogenic assays was EBM-2 media with 2.5% fetal bovine serum (Atlanta Biologicals), 100 U/mL penicillin, 100 U/mL streptomycin, and 250 ng/mL amphotericin B (Thermo Scientific).

2.3.1. HUVEC tube formation assay

150 μL of 7.8 mg/mL reduced growth factor, phenol-red free Matrigel (Corning) was polymerized in each well of a 48-well plate by incubation at 37 °C for 30 minutes. 1.2x10⁵ cells/mL were suspended in control media with or without peptide treatment. Cell solutions (200 μL) were placed on polymerized Matrigel, and incubated at 37 °C with 5% CO₂ for 8 hours before imaging. Fluorescent images of live cells were obtained (0.5 μL/mL calcein AM, Invitrogen) using a Nikon Eclipse Ti 2000 inverted light microscope equipped with a temperature/humidity controlled chamber (Pathology Devices). Images were converted to 16-bit greyscale and inverted using ImageJ [27] prior to quantification in Angioquant [28].

2.3.2. In vitro simulation of controlled release

A modified version of the HUVEC tube formation assay was developed to simulate bolus versus continuous treatment and emulate controlled release conditions. HUVECs (60–70% confluency) were either pre-treated with a high concentration (1 μM peptide or 10 ng/mL vascular endothelial growth factor, VEGF) or a low dose (0.01 μM peptide or 1 ng/mL VEGF), before being washed in PBS and resuspended in control media (0 μM) or the same low dose and seeding on Matrigel for bolus and continuous treatment, respectively (Figure 2A). Peptide doses were chosen such that dose-times were approximately equal (0.01 μM x 8 hours ≈ 1.0μM x 5 min). Cells were then imaged and tube length quantified as in section 2.3.1.

Figure 2.

In vitro assay of bolus versus continuous peptide delivery. A) Scheme of experimental setup and B) increase in HUVEC tube length upon bolus and continuous peptide treatment. A) In the bolus group, cells are pre-treated with 1 μM for 5 min, then rinsed and treated with control media for the remained of the experiment. In the continuous group, cells were treated with 0.01 μM of peptide/VEGF for both the pre-treatment period and the remainder of the experiment. Indicated concentrations are for peptides. B) Increase in tube length upon bolus and continuous treatment of HUVECs with SPARC₁₁₃(N), SPARC₁₁₈(N), and Scrambled, as well as VEGF positive control (bolus dose 10 ng/mL; continuous dose 1 ng/mL). & p<0.01, # p<0.0001 vs. within-plate control media. n=9, error bars represent standard error of the mean.

2.3.3. In vitro efficacy of degraded enzymatically-responsive hydrogels

Hydrogels were formed, incubated overnight in buffer at 37 °C, and degraded with 10 nM MMP2 (1 mL/gel). To ensure activity, MMP2 (10 μmol/mL) was supplemented every 48 hours until hydrogels had fully degraded. MMP2 was then removed from the degraded gel solution by centrifugal filtration (Ultracel 30K, Millipore), and the amount of “2T” peptide in solution determined as previously described (section 2.2.3). Degraded gel solutions were diluted in control media and used in the tube formation assay (section 2.3.1), maintaining the buffer: media ratio across all groups.

2.4. In vivo efficacy of enzymatically-responsive hydrogels

All animal procedures were approved by the University of Rochester’s University Committee of Animal Resources (UCAR).

2.4.1. Hydrogel implantation

Subcutaneous implantation was selected to evaluate in vivo vascularization as it is a well-established method to assess pro-angiogenic drugs/drug-releasing materials and allows for rapid assessment of angiogenesis after 1 week [23, 29–31]. Hydrogels were formed in silicone reactors and swollen in PBS at 37 °C overnight. 6–8 week old female BALB/c mice were obtained from Taconic (Hudson, NY). Anesthesia (60 mg/kg ketamine and 4 mg/kg xylazine) was administered via intraperitoneal injection. For pain management, buprenorphine (3.25 mg/kg) was given subcutaneously immediately before surgery and every 12 hours after as needed. Fur was removed from the back using clippers and Nair treatment. Mice were randomly assigned to treatment groups. One subcutaneous pocket was formed on each side of the mouse dorsal flank, and one hydrogel placed in each pocket with the open circular faces of the reactor facing the underlying tissue and skin. For treatments in solution (PBS, peptide in PBS (SPARC₁₁₃(N) and SPARC₁₁₈(N) at 20.6 mM, equivalent to the drug concentration in one hydrogel), or VEGF in PBS (80 μg/mL [22]; PeproTech)), reactors were placed in the subcutaneous pocket and 40 μL of solution injected into the reactor. Incision sites were closed using skin glue.

2.4.2. Fluorescent tracking of hydrogel degradation in vivo

Hydrogel persistence in vivo was quantified using the IVIS live animal imaging system as previously described [23, 32], with λ_ex/em = 570/620 nm (Xenogen IVIS-200 Optical In Vivo Imaging System, Caliper Life Sciences Inc.). Hydrogel fluorescence in terms of total radiant efficiency [(p/s)/(μW/cm²)] was quantified by selecting a ROI of constant area around each gel, and normalized to the average day 0 value for the treatment group. In vivo hydrogel degradation was fit to a pseudo-first-order degradation model using a modified version of eqn. 1, with the relative fluorescent intensity used in place of the relative hydrogel mass.

2.4.3. Assessment of vascular ingrowth

7 days after surgery, mice were sacrificed by CO₂ inhalation followed by cervical dislocation. Reactors containing residual hydrogel material and ingrown tissue were dissected and tissue homogenized by sonication in 1 mL Drabkin’s reagent (RICAA Chemical). Samples were centrifuged at 14,000 g for 20 minutes and the supernatant filtered through 0.45 μm polyvinylidene fluoride (PVDF) filters (PerkinElmer) to remove particulates. The hemoglobin concentration in each sample was determined via absorbance (A₅₄₀-A₆₅₀ nm), and comparing to a hemoglobin (Alfa Aesar) standard curve. This assessment method correlates well with other methods used to quantify vascularization [33–35].

2.4.4. Multiphoton fluorescent imaging

One week after surgery, 50 μL of 10 mg/mL 2,000 kDa fluorescein isothiocyanate-dextran (FITC, LifeTechnologies) was injected retro-orbitally into anesthetized mice. After sacrifice, the dorsal flanks were opened, and vascularization within reactors imaged in situ. Excitation light was generated using a MaiTai Ti:Sapphire laser and directed towards the sample through a BX61WI upright microscope (Olympus). An Olympus Fluoview FV300 scanning system further controlled beam scanning and image acquisition. The light directed to the scan box to image all samples was 350 mW at 810 nm, with 100 fs pulses at 80 MHz. An Olympus UMPLFL20XW water immersion lens (20×, 0.95 N.A.) was used to focus the excitation light to the sample and collect the backscattered fluorescent and second harmonic generation (SHG, collagen) signal. Note that SHG imaging can be used only for fibrillar collagens and does not allow for collagen IV imaging within vessel walls. Signals were first separated from the excitation beam using a 670 nm short pass dichroic mirror, and subsequently separated from each other using a 475 nm long pass filter. The FITC and SHG emission signals were filtered using a 535 nm (HQ535/40m-2P, Chroma) and 405 nm band-pass filters (HQ405/30m-2P, Chroma), respectively. Hamamatsu HC125-02 photomultiplier tubes were used to collected signals from both channels. Images were collected at the same five pre-determined locations within each reactor. At each location a 100 μm stack was taken as 21 images spaced 5 μm apart. Following image collection stacks were maximum intensity projected and the vessel diameters measured manually in ImageJ [27]. Five of the largest vessels were measured per image, giving 25 measurements (e.g., five vessels each from five locations) that were subsequently averaged to give a single average vessel diameter for each gel. Image collection and vessel measurements were performed by a user blinded to treatment groups.

2.5. Statistical analysis

Data was assembled and calculations were formed in Microsoft Excel (2010 v14.0). Statistical analysis and figure preparation was performed using Graphpad Prism (5.04). Data is presented as mean ± standard error of the mean for all figures and reported values. Data in Figure 3 (Scrambled peptide), 5, 7, and diffusivity coefficients were analyzed using a one-way ANOVA, with post-hoc testing as appropriate (Figure 5 with Dunnett’s and Figure 7A with Tukey’s post-hoc testing). Data in Figure 2, 3 (SPARC₁₁₃ and SPARC₁₁₈), 4, 6, and Table 2 were analyzed using a two-way ANOVA, with post-hoc testing as appropriate (Figure 2, Figure 3 (SPARC₁₁₃ and SPARC₁₁₈), Figure 4, and Table 2 with Bonferroni post-hoc testing). All experiments were performed at least in duplicate to reach n=6–16, as specified in figure legends. p<α=0.05 was considered significant for all analyses.

Figure 3.

Dose response curves for “N” (open symbols) and “2T” (solid symbols) peptides. A) Representative images (at 1 μM) and quantification of tube length upon treatment with B) SPARC₁₁₃, diamonds and C) SPARC₁₁₈, circles. The scrambled peptide (grey squares) did not affect tube formation at any concentration. * p<0.05, & p<0.01, $ p<0.001, # p<0.0001 vs. 0 μM (control media) treatment. n=6–9; error bars represent standard error of the mean. Scale bar = 250 μm.

Figure 5.

Dose response curves for degraded hydrogel products, in terms of fraction of a gel per well. A) Representative images at 1/70^th of a gel/well and quantification of tube length upon treatment with B) SPARC₁₁₃-releasing gels, diamonds and C) SPARC₁₁₈-releasing gels, circles. The scrambled peptide releasing gels (squares) did not affect tube formation at any concentration. * p<0.05, & p<0.01, $ p<0.001, # p<0.0001 vs. 0 μM (control media) treatment. n=9; error bars represent standard error of the mean. Scale bar = 250 μm.

Figure 7.

Vascularization of hydrogels one week after implantation. Hydrogels releasing SPARC₁₁₃ and SPARC₁₁₈ in response to MMPs were formed in non-degradable reactors and implanted subcutaneously, and compared to gels releasing scrambled peptide, free SPARC₁₁₃ and SPARC₁₁₈ in solution, and PBS and VEGF controls. After one week, vascularization was assessed via A) hemoglobin content within the reactor and B–C) by imaging vasculature formed using multiphoton fluorescence imaging. B) Quantification of average vessel diameter. C) Representative images of vasculature (FITC-dextran; red) and collagen (SHG; blue). n.s. p>0.05, * p<0.05, & p<0.01, $ p<0.001, # p<0.0001. n=16 for A, 6–8 for C; error bars represent SEM. Scale bar = 100 μm.

Table 2.

Summary of experimentally-determined degradation kinetic time constants (k_deg).

Enzymatically-responsive hydrogel	In vitro kdeg (dry mass, with 10 nM MMP2) (x10−2; hour−1)	In vivo kdeg (fluorescence intensity) (x10−2; hour−1)
SPARC113(DL)	1.16 ± 0.03b,c	1.46 ± 0.12
SPARC118(DL)	8.77 ± 0.74a,c,2	1.44 ± 0.181
Scrambled(DL)	3.23 ± 0.13a,b,2	1.50 ± 0.241

Within degradation environment (columns), across gel type (rows):

^ap<0.05 vs. SPARC₁₁₃(DL),

^bp<0.05 vs. SPARC₁₁₈(DL),

^cp<0.05 vs. Scrambled(DL).

Within gel type (rows), across degradation environment (columns):

¹p<0.05 vs. in vitro,

²p<0.05 vs. in vivo.

n=6 (in vitro) or 16 (in vivo); mean ± standard error of the mean.

Figure 4.

In vitro degradation and peptide release from enzymatically-responsive hydrogels. (i) Mass loss, (ii) swelling ratio, (iii) “2T” peptide release from enzymatically responsive hydrogels, and (iv) diffusive release of encapsulated peptide from non-degradable gels. Gels were incubated in buffer alone (solid symbol and line) for 24 hours, at which point 10 nM MMP2 was added to a subset of gels (open symbol, dashed line). A) SPARC₁₁₃(DL), B) SPARC₁₁₈(DL), and C) Scrambled(DL) gels were investigated. * p<0.05, & p<0.01, $ p<0.001, # p<0.0001 versus gel in buffer alone at same time point. n=6 except for the swelling data where gel degradation reduced sample size at later time points; error bars represent standard error of the mean (some are obscured by symbol).

3. Results and discussion

Exploiting our previously developed platform technology [15], this study rigorously investigated the therapeutic potential of PEG hydrogels providing enzymatically-responsive release of pro-angiogenic peptides. Previous work identified three bioactive peptide releasing hydrogels: Qk(DL), SPARC₁₁₃(DL), and SPARC₁₁₈(DL). However, only SPARC₁₁₃(DL) and SPARC₁₁₈(DL), and hydrogels releasing a scrambled control peptide, Scrambled(DL), were investigated here. Hydrogels releasing Qk, a VEGF mimic, were omitted due to comparably slow degradation and poor peptide release (for more information see [15]). The efficacy of continuous versus bolus delivery of SPARC-based peptide drugs was investigated. Hydrogel degradation, peptide release behaviors, and pro-angiogenic efficacy of the drug delivery system was characterized in vitro and in vivo.

3.1. SPARC peptides benefit from continuous delivery and show in vitro activity at therapeutically-relevant concentrations

Many drugs have been shown to be more or only efficacious under controlled/sustained release conditions [4, 14, 22]. However, to the best of our knowledge, the same has not been shown for the SPARC peptides studied here. Therefore, a commonly used in vitro measure of angiogenic potential was modified to simulate bolus versus continuous delivery (Figure 2A). While unable to induce tube formation upon bolus treatment (1.2-fold control media), continuous treatment with a lower concentration of VEGF significantly increased tube length by 1.8-fold. These results were expected, as activation of the VEGF signaling pathway (VEGF/VEGFR2 interactions leading to MAPK and Akt phosphorylation) is known to depend on dose and duration of VEGF treatment [36, 37], and VEGF has been show to benefit from controlled release in vivo [4, 22].

Similar to VEGF, neither SPARC₁₁₃(N) nor SPARC₁₁₈(N) treatment resulted in significant tube network formation upon bolus delivery (1.1 and 1.3-fold control media, respectively). However, both were bioactive with continuous delivery (2.7 and 1.8-fold versus control media for SPARC₁₁₃(N) and SPARC₁₁₈(N)). While the signaling pathways for the SPARC peptides are not well understood, GHK, common to both peptides, has been shown to activate the angiotensin (AT) II AT1 receptor [38]. Activation of the AT1 receptor, similar to VEGF/VEGFR2 activation, has been shown to depend on therapeutic dose and treatment duration [39], supporting our findings that sustained delivery of the SPARC peptides is necessary for tube formation, motivating the incorporation and controlled release of these peptides from hydrogel depots.

Dose-response curves of the drug alone (native “N”) and expected released (two-tailed “2T”) forms of the SPARC peptides were investigated using the HUVEC tube formation assay (Figure 3) to identify likely therapeutic concentrations for successful in vivo translation.

Both SPARC peptides in “N” and “2T” forms increased the formation of smooth, honeycomb-like tube networks as compared to scrambled peptide or media alone. The scrambled peptide did not significantly affect tube length at any concentration, indicating that the observed results are sequence-specific. SPARC₁₁₃ significantly increased tube length at 0.01, 0.1, 1, and 10 μM (2.5, 2.8, 3.0, and 2.5-fold, respectively) in its “N” form, but only at 100 μM in the “2T” form (2.6-fold, Figure 3B). Similarly, SPARC₁₁₈ significantly increased tube length at all explored concentrations in its “N” form (1.8, 2.0, 1.9, 2.0, and 2.2-fold at 0.01, 0.1, 1, 10, and 100 μM, respectively), but only at 0.01 μM in the “2T” form (1.9-fold, Figure 3C). Tube network formation with both SPARC peptides was significantly affected by concentration and the presence of the amino acid “tails”, with SPARC₁₁₃ demonstrating a significant interaction between these factors, while SPARC₁₁₈ did not (Supplemental Table 1). These results indicate that residual MMP substrates (2T) adversely affect, but do not abrogate, SPARC peptide bioactivity. Taken together, these results suggest that both SPARC₁₁₃(DL) and SPARC₁₁₈(DL) gels could provide local therapeutic levels of peptide, as one 40 μL gel contains ~ 0.8 μmol of drug. Furthermore, SPARC₁₁₃-releasing gels are likely more promising due to the steadily increasing tube formation that occurs upon exposure to increasing doses of SPARC₁₁₃(2T).

SPARC₁₁₃ has previously been shown to increase vessel formation in the chicken chorioallantoic membrane (CAM) assay at 10–500 μM, with a maximum increase in capillary density occurring at 50 μM [17]. The observed biphasic concentration behavior is consistent with our results. SPARC₁₁₉ (KGHK rather than SPARC₁₁₈ which is KKGHK) causes dose-dependent increases in capillary density in the CAM assay as peptide concentration is increased from 10 to 5,000 μM [17]. However, less dramatic dose-dependency was observed here with respect to tube length upon treatment with SPARC₁₁₈(N).

3.2. Enzymatically-responsive hydrogels degrade and release bioactive peptides

SPARC₁₁₃(DL), SPARC₁₁₈(DL), and Scrambled(DL) hydrogels were formed, swollen overnight, and then degraded via treatment with supraphysiologic concentrations of MMP2 [40]. Gels degraded and released peptide in the presence of MMP2, but were stable in buffer alone (Figure 4), illustrating enzymatically-responsive behaviors. Specifically, SPARC₁₁₃(DL), SPARC₁₁₈(DL), and Scrambled(DL) gels degraded in the presence of the enzyme over 144, 30, and 72 hours, respectively. This is consistent with previous work showing that central peptide size and hydrophobicity affects hydrogel degradation behavior [15]. SPARC₁₁₃(DL), SPARC₁₁₈(DL), and Scrambled(DL) gels swelled to different degrees after 24 hours in buffer (13, 21, and 11 mg/mg, respectively), but all degraded via bulk mechanisms as indicated by the significant increase in swelling ratio over time for MMP-treated gels, consistent with expectations as gel mesh sizes are significantly larger than MMP2 (19–45 nm calculated as in [41], compared to ~ 2.6 nm for MMP2 [42]).

Peptide release kinetics from degrading gels was in close agreement with hydrogel degradation kinetics. SPARC₁₁₃(DL), SPARC₁₁₈(DL), and Scrambled(DL) gels in MMP2 released 8, 26, and 22% of peptide upon reverse gelation, respectively. In contrast, gels incubated in buffer did not release peptide. When encapsulated within, rather than covalently linked to, non-degradable gels, all three peptides were rapidly released (80–98% of encapsulated peptide released over 3 hours). There were no significant differences in diffusion coefficients for the three peptides (p>0.05), with rates ranging from 0.7 to 1.4 x10⁻¹⁰ m²/sec [26]. These results demonstrate the importance of using the enzymatically-responsive tether to extend peptide release. They also confirm that the differences in peptide release profile observed are a result of differences in the rate of “DL” cleavage and liberation of peptide from the hydrogel, and not due to differences in diffusion rates out of gel networks.

Degraded gel solutions containing released peptides were assessed for bioactivity using HUVEC tube formation assays. As gels release varying amounts of peptide in “2T” form, gel/well ratios were chosen to compare bioactivity of the peptide-releasing hydrogels. Thus, total drug concentration between groups is kept constant and the assay better represents how peptides will be liberated in vivo. 1/70 of a gel/well was selected as the highest concentration as it corresponds to 10 μM of SPARC₁₁₈(2T), near the upper limit of the doses investigated here (Figure 3). However, variations in the amount of peptide fully released from the PEG macromers resulted in different released “2T” peptide doses, with 1/70 of a gel/well corresponding to 13 and 24 μM of SPARC₁₁₃(2T) and Scrambled(2T), respectively. Serial log dilutions were used to investigate dose-dependent behavior on tube formation.

Degraded SPARC₁₁₃(DL) hydrogels significantly increased HUVEC tube length to 1.6, 1.7, 1.7, and 2.1-fold that of control media at 1/70,000, 1/7,000, 1/700, and 1/70 of a gel/well, respectively (Figure 5). While degraded SPARC₁₁₈(DL) hydrogels resulted in trending increases in HUVEC tube length at all concentrations investigated (3.1, 2.8, and 2.8-fold at 1/70,000, 1/7,000, and 1/700 of a gel/well, respectively), only the 4.3-fold increase at 1/70 of a gel/well was statistically significant. The degraded Scrambled(DL) hydrogels did not significantly affect tube formation at any concentration investigated. This demonstrates that SPARC₁₁₃(DL) and SPARC₁₁₈(DL) hydrogels release sequence-dependent bioactive components. SPARC₁₁₈(DL) hydrogels resulted in more substantial increases in HUVEC tube length than SPARC₁₁₃(DL) gels at every dose investigated (e.g., 4.3 vs. 2.1-fold at 1/70 of a gel/well). However, the SPARC₁₁₈(DL) gels also had substantially more intra-group variability and only significantly increased tube length at the highest dose investigated. Overall, the SPARC₁₁₃(DL) gels showed much more robust proangiogenic behavior versus SPARC₁₁₈(DL), as they resulted in significant, albeit less dramatic, increases in tube length over a 1,000-fold range of concentrations.

Tube formation due to a range of released peptide doses are shown in Supplemental Figure 1. Together with Figure 3, these data highlight that the efficacy of the SPARC₁₁₃(DL) hydrogels cannot be fully attributed to the bioactivity of free drug alone. Degraded hydrogels significantly increased tube length even when the amount of “2T” present in solution is below the minimum effective concentration for “2T” peptide alone. For example, SPARC₁₁₃(DL) at 1/700 of a gel/well significantly increases tube length. However, there is only 1.3 μM of SPARC₁₁₃(2T) in the 1/700 of a gel/well solution, and free SPARC₁₁₃(2T) only significantly increases tube length at 100 μM. This indicates that SPARC₁₁₃ remaining tethered to PEG retains bioactivity. This is consistent with previous work that has shown that PEG conjugation (PEGylation) does not preclude growth factor bioactivity [9, 43]. It also suggests the possibility that tethering SPARC₁₁₃ to PEG enhances bioactivity, potentially by increasing peptide co-localization at receptors, although additional studies would be necessary to confirm this possibility.

3.3. Enzymatically-responsive hydrogels degradation rate is altered between in vitro and in vivo environments

Hydrogels that included fluorescently-labeled peptides (FLP*) were implanted subcutaneously and degradation tracked longitudinally using live animal imaging [44]). All three hydrogels degraded in vivo over 1 week (Figure 6), with fluorescence approaching baseline levels (PBS controls) after ~ 4 days (Supplemental Figure 3). Scrambled(DL) hydrogels degraded slightly more slowly than drug-releasing hydrogels, with degradation significantly affected by time and hydrogel type (p<0.05), but without interaction between these factors (p>0.05).

Figure 6.

Hydrogel degradation in vivo. A fluorescently-tagged peptide (FLP*) was covalently linked to the hydrogels and gel fluorescent tracked using the IVIS live animal imaging system. In vitro testing confirmed that longitudinal hydrogel fluorescence correlates well with degradation (Supplemental Figure 2). A) Representative fluorescent images and B) quantification of fluorescence over the one week study, normalized to average within-group day 0 fluorescence. Gel degradation is significantly affected (p<0.0001) by gel type and time, but there was no interaction between the factors (p>0.05). n=16; error bars represent standard error of the mean.

Kinetic time constants (k_deg) were calculated to describe the rate of hydrogel degradation in vitro and in vivo based on data in Figures 4 and 6. As shown in Table 2, in the presence of supraphysiological concentrations of MMP2 (10 nM in vitro, compared to ~ 0.15 pM in vivo [40]), SPARC₁₁₈(DL) hydrogels had the highest k_deg, significantly greater than Scrambled(DL), which was significantly higher than the SPARC₁₁₃(DL) hydrogels. Kinetic time constants were not calculated for hydrogels in buffer alone, as no degradation was observed (Figure 4). In vivo there were no significant differences in k_deg between the three gel types. SPARC₁₁₃(DL) had statistically equivalent time constants in vitro and in vivo; however, both SPARC₁₁₈(DL) and Scrambled(DL) gels had significantly lower kinetic time constants in vivo, indicating slower degradation.

Differential expression of MMPs occurs in wounded tissue, with the tissue, type of injury, and time after injury affecting levels and specific MMPs expressed [40, 45, 46]. For example, basal levels of active MMP2 and 9 in murine hind limbs are ~ 0.15 and ~ 0.6 pM, respectively. Upon introduction of ischemia by femoral artery ligation, these levels increase ~ 10-fold, with maximal MMP2 activity occurring 14 days after surgery, while maximal MMP9 activity occurs after 3–7 days [40]. In contrast, maximal MMP1 (6.5-fold control) and MMP2 expression (6-fold control) in cardiac tissue occur 7 days after onset of ischemia via coronary artery ligation, while MMP9 expression is highest (4-fold control) after 4 days [46]. As the degradable sequence used here, “DL”, is susceptible to cleavage by numerous MMPs (1, 2, 3, 7, 9, 14) [47], additional MMPs likely contributed to hydrogel degradation. MMP expression is likely increased subcutaneously as a result of local tissue injury due to hydrogel implantation, although likely not to similar levels as in these ischemic models, as subcutaneous implantation represents a much less severe injury. This is highlighted by SPARC₁₁₃(DL) hydrogel degradation rate equivalence in vitro and in vivo, despite supraphysiological MMP2 concentrations used in vitro [40]. However, both the SPARC₁₁₈(DL) and Scrambled(DL) hydrogels degraded more slowly in vivo than in vitro. This result underlines an inherent limitation of the in vitro degradation model: while indicative of the enzymatically-responsive nature of the gels, it does not predict degradation rates in vivo. The slowed in vivo degradation for SPARC₁₁₈(DL) and Scrambled(DL) hydrogels implies two possibilities: either these gels are less susceptible to cleavage by the additional MMPs present subcutaneously than the SPARC₁₁₃(DL) hydrogels, and/or the peptide drugs affect MMP production by the host tissue, with fewer or lower levels of MMPs expressed in response to the SPARC₁₁₈(DL) and Scrambled(DL) hydrogels than the SPARC₁₁₃(DL) gels. GHK-containing peptides have been shown to affect MMP expression by fibroblasts in vitro [48] and in cutaneous wounds in vivo [49], supporting the theory that SPARC peptide-releasing gels may affect MMP expression. However, the analysis conducted here is unable to determine if either or both of these factors contributes to the observed differences in degradation kinetics. Additional analysis such as zymography could be employed to determine the relative levels of MMPs expressed in vivo. This data, coupled with degradation and peptide release studies performed with additional MMPs would be necessary to address this question. Physiologically-relevant degradation solutions that better emulate in vivo MMP expression, or co-culture of hydrogels with cells (or ex vivo tissue samples) stimulated with pro-inflammatory factors such as tumor necrosis factor-alpha [50], could be used to more accurately predict in vivo gel degradation and peptide release profiles. It is also possible that in vivo hydrogel degradation was affected by aspects of the in vivo environment beyond MMP expression such as accelerated hydrolytic degradation of the gels [51] by macrophages recruited during the inflammatory response to hydrogel implantation [52].

3.4. Both SPARC-releasing hydrogels induce vascularization in vivo

The extent of vascular ingrowth within pro-angiogenic hydrogels was assessed by quantifying hemoglobin (Hb) content, after confirming that hydrogel fluorescent labeling does not affect Hb quantification (Supplemental Figure 4). SPARC₁₁₃(DL) hydrogels resulted in the greatest extent of vascular ingrowth (745 μg Hb/reactor), significantly more than Scrambled(DL) and PBS negative controls (388 and 430 μg Hb/reactor) and free SPARC₁₁₃(N) (219 μg Hb/reactor; Figure 7). This corresponds to 1.7, 1.9, and 2.5-fold increases over PBS, scrambled peptide releasing gels, and SPARC₁₁₃(N), respectively. SPARC₁₁₈(DL) hydrogels caused similar increases in vascular ingrowth (633 μg Hb/reactor), corresponding to a significant 1.6-fold increase in vascular ingrowth over scrambled peptide releasing gels, a significant 2.0-fold increase in vascular ingrowth compared to free SPARC₁₁₈(N) (310 μg Hb/reactor), and an insignificant 1.5-fold increase over PBS. There was no significant difference in Hb levels between SPARC₁₁₃(DL) and SPARC₁₁₈(DL) hydrogels. Bolus delivery of SPARC₁₁₃(N) and SPARC₁₁₈(N) in PBS did not result in angiogenesis (both 0.7-fold PBS), while the positive control VEGF caused trending, although statistically insignificant, increases in vascular ingrowth (612 μg Hb/reactor, 1.4-fold PBS). A summary of all statistical comparisons is shown in Supplemental Table 2.

Multiphoton fluorescence imaging was used to qualitatively assess vascularization due to pro-angiogenic peptide-releasing hydrogels in situ, without additional tissue manipulation and processing [53]. While the average vessel diameter within the reactor varied from 17 (VEGF) to 28 μm (SPARC₁₁₈(N) and Scrambled(DL)), there were no significant differences between the groups. Interestingly, vessel size exhibited opposite trends from the Hb data, with treatments that resulted in more Hb/reactor (e.g., SPARC₁₁₈(DL)) trending towards smaller vessels than those with less Hb/reactor (e.g., SPARC₁₁₈(N)). Taken together, this data, as well as the representative images shown in Figure 7C, supports the conclusion that controlled SPARC₁₁₃(DL) and SPARC₁₁₈(DL) delivery via MMP-mediated liberation from hydrogels resulted in the formation of smaller, but more plentiful, vasculature than the other treatment groups. This presumably provided better perfusion of the hydrogels, thereby increasing hemoglobin level. These results demonstrate the ability of the SPARC₁₁₃ and SPARC₁₁₈-releasing hydrogels to enhance angiogenesis, and illustrate the importance of controlled release for pro-angiogenic therapies. Similar to our findings, PEG hydrogels that exhibit sustained release of VEGF via enzymatic activity significantly increased implant vascular volume and reperfusion of ischemic hindlimb tissue [22]. Conjugation of VEGF to enzymatically-degradable PEG hydrogels increased endothelial cell proliferation [54] and resulted in localized neovascularization using the CAM assay [43]. The GHK peptide sequence within SPARC₁₁₃ and SPARC₁₁₈ has been shown to increase fibroblast growth factor (FGF) and VEGF expression by fibroblasts [55] and mesenchymal stem cells [56]. Therefore, it is possible that the peptide-releasing hydrogels increase pro-angiogenic growth factor expression of local tissue, which could contribute to the observed vascular response.

Based on promising results presented here, it can be concluded that the ~ 4 days of peptide delivery achieved from the SPARC₁₁₃(DL) and SPARC₁₁₈(DL) hydrogels is sufficient to enhance angiogenesis in vivo. However, this is much more rapid than many pro-angiogenic protein delivery systems, which often deliver proteins over ~ 2–4 weeks [4, 22, 57]. As the duration of pro-angiogenic factor delivery is critical for vessel formation [4], it is possible that this temporal release profile, while sufficient, is not ideal for SPARC₁₁₃ and SPARC₁₁₈, and that altering the duration of delivery could further increase vascularization. As ischemia has been shown to affect MMP expression [40, 45, 46], it is likely that hydrogels would degrade over differential, and not necessarily optimal, time scales in ischemic tissue. The time over which degradation and peptide release occurs could be manipulated by changing the degradable linkage used [23, 24, 47]; however, this approach has the limitation of altering the “tails” on the peptide drug, likely affecting peptide bioactivity [15]. Alternately, hydrogel crosslinking density could be increased [58]; however, this would also affect hydrogel drug capacity. Use of the non-invasive tracking system utilized here, combined with these methods to alter hydrogel degradation time scale, could identify optimal temporal delivery profiles of peptide drugs for specific target tissues/disease states.

The therapeutic effects of these hydrogels is not limited to vascularization applications: the sequence GHK contained within both SPARC peptides has numerous beneficial effects not assessed in this study, including anti-inflammatory effects, improving cell recovery after X-ray damage, increasing extracellular matrix protein synthesis, and improving nerve outgrowth [18]. GHK also increases the production of pro-angiogenic factors by mesenchymal stem cells (MSCs) [56], suggesting that these hydrogels may improve the therapeutic efficacy of encapsulated or nearby host MSCs [59].

4. Conclusions

Overall, the materials exploited here for the controlled delivery of pro-angiogenic peptides hold great therapeutic potential: both the SPARC₁₁₃ and SPARC₁₁₈-releasing hydrogels resulted in significant angiogenesis upon subcutaneous implantation, showing potential to treat ischemic tissue disorders or promote vascularization within tissue engineering approaches. While these results are promising, additional studies are required in appropriate diseased tissue and/or tissue engineering models to fully investigate pro-angiogenic hydrogel therapeutic potential. Additionally, both gels hold promise in other wound healing and cell-based tissue engineering approaches, as they contain the GHK sequence shown to affect numerous aspects of wound repair and cellular behavior.