Two experimental designs were utilized:

• Assessment of the model's sensitivity to recipient sites containing different vascular densities (Table 1)

• Determination of whether the model is permissive to identifying changes in vascular density using VEGF-loaded microspheres (Table 2).

Specific procedures and details for the experimental groups listed in Tables 1 and 2 follow.

Figure 1

Table 1: In vivo implantation sites

Figure 2

Table 2: Application of VEGF microspheres

The model TEC's design was based, in part, on an initial concept of Dvorak et al.₉ The TEC shell was constructed from polypropylene film using a matched die mold in a heated horizontal press. The TEC is designed to be radially symmetric to enable future mathematical modeling. The utilization of polypropylene as the shell limits the invasion of tissue into a single surface of the TEC. The internal TEC diameter measures 10.5 mm, tapering to 8.5 mm over a height of 4 mm (Fig. 1). The integral sewing ring extends off of the 10.5 mm diameter end for an additional 3.5 mm, making the total diameter 17 mm. The TECs were sterilized with 100% ethanol and allowed to air dry prior to applying fibrin gels. Fibrin was selected as an initial single component ECM for this study. On the day of implantation, each TEC was filled with Tisseel® Fibrin Sealant (Baxter-Hyland, Glendale, CA) using sterile technique under a tissue culture hood. The Tisseel® system employs a duplo-syringe, one side containing standardized fibrinogen and the other containing standardized thrombin and CaCl2. For TECs containing endogenous VEGF (rhVEGF165, R&D Systems, Minneapolis, MN), the growth factor was added as an aqueous solution to the fibrinogen syringe of the Tisseel® system prior to injecting the fibrin into the TEC. The final VEGF concentration was 5 ng/mL.

Figure 3

Fig. 1: Schematic diagram of the model TEC.

Microsphere manufacture using the solid encapsulation/single emulsion/solvent extraction technique and VEGF loading was performed as previously described in detail.₁₀,₁₁ The fabrication protocol results in biodegradable polymer microspheres that contain numerous microdomains of encapsulated VEGF (see Movie).

Microspheres containing rhVEGF165 (R&D Systems, Minneapolis, MN) were manufactured by co-encapsulating the VEGF with rat serum albumin (RSA) as a carrier protein. The ratio of VEGF:RSA (w/w) was 1:2,000. The VEGF-loaded microspheres release 70 ng/mL of VEGF at 4 days, followed by a steady state VEGF concentration =5 ng/mL for at least 28 days.¹⁰ For the VEGF-loaded microsphere study, all TECs were implanted on skeletal muscle.

TEC implantation was conducted as previously described.₁₂ To determine the model's sensitivity in assessing vascular density ( v), TEC implants were placed in two different anatomical sites (dermis and skeletal muscle) that possess different v and global capillary direction. Skeletal muscle possesses a higher v than dermis in the rat. Moreover, capillaries in the skeletal muscle are largely in the direction of muscle fibers, that being tangent to, rather than perpendicular to, the TEC's open surface. A longitudinal skin incision was made along the spine and using blunt dissection, the skin was separated from the underlying tissue. For implants on the dermis, the panniculus carnosus (cutaneous trunci) muscle was removed and, if necessary, pressure was used to achieve hemostasis. For TEC implantation on skeletal muscle, a uniform muscle surface on the back was exposed and the thin fascia covering the muscle body was removed. The TEC was sutured directly onto the exposed uniform muscle body (rather than multiple muscle bellies) as described above (Fig. 2). Each rat received four TECs, one from each experimental group shown above in Table 1. Four animals per time point were selected based upon preliminary experimental evaluation and power-law analysis (data not shown).

Figure 4

Fig. 2: Perioperative view of fibrin-filled TEC sutured to skeletal muscle.

Animal euthanasia and harvest were conducted as previously described.?? Post-implantation days ranging from 5 to 14 days were selected as timepoints for TEC explantation. The selection of time points for explantation was based on previous wound healing investigations demonstrating capillary extension in 3-4 days and quiescence by 21 days.⁹,₁₃ Thus, explantation timepoints of 5-14 days were selected based on expected rates of neovascularization and the degradation rates of the fibrin.₁₄,₁₅

The CD31 staining protocol, digital imaging and microscopy, and quantitative assessment of vascular density were conducted as previously described in detail.¹²,₁₆ For each tissue section, five equally-spaced images were obtained along the interface of the implanted TEC and adjacent tissue surface (either dermis or muscle, Fig. 3). The underlying native tissue was not included in the image. The two lateral images of the TEC were labeled “edge”, the central image was labeled “center”, and the remaining two internal images were labeled “mid”. The total area of CD31 positive vasculature was calculated for each TEC image (total of 5 TEC) using image analysis software and a previously reported semi-automated threshholding technique.¹²,¹⁶ The v is defined as the ratio of the mean of the CD31 positive area to the total TEC area.

Data are presented in two formats: (1) the pooled v for each TEC (i.e. the mean of all analyses for each TEC regardless of geometric location within the TEC) and, (2) the v stratified to the geometric location within the TEC. Results are expressed as the mean ± SEM for four experiments. Statistical analysis was performed by the student t-test and one-way ANOVA with Newman-Keuls multiple comparison post hoc test. Statistical significance was defined as p = 0.05.

Figure 5

Fig. 3: Cross-section of explanted fibrin-filled TEC (outlined) on muscle (tissue left of outline) depicting the areas of histometric analysis.

The TECs were intact and none of the implantation sites demonstrated hematoma or seroma at the time of explantation. Furthermore, all of the TECs were adherent to their respective implantation site at the time of explant. That is, there was no observed void space between the TEC and site of implantation. In addition, the surrounding tissues showed no signs of inflammation, infection, or gross atypia. The fibrin matrix within the TECs was present at all experimental timepoints. However, the volume of matrix present diminished with respect to time.

Two implantation sites were compared in this investigation. It was hypothesized that there would be a difference in the TEC v as a function of the implantation site. The skin implantation site refers to the inferior dermal surface of skin overlying the medial back of the animal model. This was compared to implantation on skeletal muscle adjacent to the dermal implantation on the back. After 7 days of implantation there was a significantly greater total v in the VEGF(-)/Fibrin TECs implanted on the skin when compared to VEGF(-)/Fibrin TECs implanted on the muscle. However, after 14 days of implantation, no significant difference in total v was observed when comparing experimental groups based on implantation site (i.e. VEGF(+)/Fibrin-skin vs. VEGF(+)/Fibrin-muscle and VEGF(-)/Fibrin-skin vs. VEGF(-)/Fibrin-muscle; Fig. 4). Comparison of total v between VEGF(+)/Fibrin TECs and VEGF(-)/Fibrin TECs demonstrated increases in v (Fig. 4). The skeletal muscle implants demonstrated significantly increased v at 7 and 14 days, and the skin implants demonstrated significantly increased v by day 14. Temporally (i.e. comparing day 7 to day 14) there was a significant increase in the total v of the VEGF(-)/Fibrin TECs implanted on muscle (Fig. 4, bottom).

Figure 6

Fig. 4: Angiogenesis expressed by v within the TEC. (A) Day 7, (B) Day 14. The data are represented as mean ± SEM. * and ** denote a p < 0.05 and a p < 0.01, respectively, for VEGF(+)/Fibrin vs. VEGF(-)/Fibrin groups of the same anatomical location. *** denotes a p < 0.05 for VEGF(-)/Fibrin-skin vs. VEGF(-)/Fibrin-muscle. NC denotes no change in the v comparing day 7 to day 14. The vertical arrow denotes a significant increase in v comparing day 7 to day 14.

During the course of image analysis, trends in location of vascularization were noted. In Figs. 5 and 6, the total v have been stratified to location within the TEC. Figs. 5A and 5B represent the relative v of TECs implanted on skin at 7 and 14 days, respectively. Initially, at 7 days of implantation, the v at the edge of the TEC was significantly increased in the VEGF(+)/Fibrin TECs when compared to VEGF(-)/Fibrin TECs. However, by 14 days of implantation, the central region of the VEGF(+)/Fibrin TECs had developed a significantly greater v when compared to VEGF(-)/Fibrin TECs. Figs. 6A and 6B demonstrates similar trends at the skeletal muscle implantation site. In this case, VEGF(+)/Fibrin TECs demonstrated a significant increase in v at the edge of the TEC at day 7 and a significant increase in v in the central region at day 14 when compared to VEGF(-)/Fibrin TECs. Temporally, there was a significant increase in v at the edge of the TEC for fibrin implants on muscle when comparing day 7 to day 14.

Figure 7

Fig. 5: Angiogenesis expressed by v with respect to position within the TEC for skin implants at (A) day 7 and (B) day 14. The data are represented as mean ± SEM. * denotes a p < 0.05 for VEGF(+)/Fibrin vs. VEGF(-)/Fibrin groups of the same TEC location. The cartoon represents the location of the data within the TEC. NC denotes no change in the v comparing day 7 to day 14.

Figure 8

Fig. 6: Angiogenesis expressed by v with respect to position within the TEC for muscle implants at (A) day 7 and (B) day 14. The data are represented as mean ± SEM. * denotes a p < 0.05 for VEGF(+)/Fibrin vs. VEGF(-)/Fibrin groups of the same TEC location. The cartoon represents the location of the data within the TEC. NC denotes no change in the v comparing day 7 to day 14. The vertical arrow denotes a significant increase in v comparing day 7 to day 14.

Overall, total vascular density within all experimental groups was greater than 5% for all timepoints (data not shown). Comparatively, vascular density of rat skeletal muscle averaged 2-3% (data not shown). Stratifying the data to physical location within the TEC reveals no statistical difference in vascular density with respect to physical location within the TEC at Day 5 (see Fig. 7A-C). By day 10, the VEGF microspheres had a significantly greater vascular density at both the center and edge of the TEC (see Fig. 7D-F). At the center of the TEC, the VEGF microspheres had significantly greater vascular density as compared to all other groups. The RSA microspheres had a significantly greater vascular density at the center when compared to empty microspheres and VEGF protein. In addition, both the VEGF microspheres and VEGF protein had a significantly greater vascular density at the edge when compared to RSA microspheres. However, by day 14 there is no statistical difference in vascular density with respect to physical location within the TEC (see Fig. 7G-I).

Figure 9

Fig. 7: Vascular Density of TECs With Respect to TEC Geometry and Time. The legend refers to the type of TEC implanted. That is, Empty µs = TECs containing empty microspheres; RSA µs = TECs containing RSA microspheres; VEGF µs = TECs containing VEGF microspheres; VEGF = TECs containing VEGF protein. Data are mean ± SEM. ** represents a p < 0.01 and * represents a p < 0.05 as derived from ANOVA analysis.

Comparisons of vascular density changes within each experimental group with respect to time of implantation are presented in Fig. 8. A significant increase in vascular density was observed in the center of VEGF-encapsulated microsphere TECs between the 5 day and 10 day timepoints. There was no statistical difference between TECs filled with fibrin alone and TECs filled with fibrin + empty microspheres.

Figure 10

Fig. 8: Vascular Density Within TECs Over Time. For TEC location (A) Mid, (B) Edge, (C) and Center. Note the increased relative rate of angiogenesis at the center of the TEC for the VEGF microspheres between day 5 and 10. * represents a p <0.05as described by ANOVA analysis.

This work was supported, in part, by a Whitaker Foundation Grant (CWP), National Institutes of Health grant, HL62341 (CWP), Army DOD grant, DAMD17-99-1-9268 (CWP), and The University of Texas M.D. Anderson Cancer Center core grant from the National Cancer Institute, CA-16672.