Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2012 Nov 1.
Published in final edited form as: J Vasc Surg. 2011 Aug 27;54(5):1451–1460. doi: 10.1016/j.jvs.2011.05.050

The influence of early phase remodeling events on the biomechanical properties of engineered vascular tissues

Zehra Tosun 1, Carolina Villegas Montoya 2, Peter S McFetridge 1,*
PMCID: PMC3202686  NIHMSID: NIHMS321464  PMID: 21872418

Abstract

Objectives

Over the last decade the use of ex vivo-derived materials designed for use as implant scaffolds has increased significantly. This is particularly so in the area of regenerative medicine, or tissue engineering, where the natural chemical and biomechanical properties has been shown to be advantageous. By focusing on detailed events that occur during early phase remodeling processes our objective was to detail progressive changes in graft biomechanics to further our understanding of these processes.

Methods

Using perfusion bioreactor system and acellular human umbilical veins (HUV) as a model 3D vascular scaffold, human myofibroblasts were seeded and cultured under either static or defined pulsatile conditions. Cell function in relation to graft mechanical properties were assessed.

Results

Cells were shown to have doubled in density from approximately 1 × 106 to 2 × 106 ± 0.4 × 106 cells/cm ringlet while static cultures remained unchanged. In both static and dynamic systems the materials compressive stiffness and ultimate tensile strength remained unchanged. However the Young’s modulus values increased significantly in the physiological range while in the failure range a significant reduction (66%) was shown under dynamic conditions.

Conclusions

We have shown that as pulse and flow conditions are modulated, complex mechanical changes are occurring that modifies the elastic modulus differentially in both physiological and failure ranges. It is clear that mechanical properties play an important role in graft patency, and that a dynamic relationship between structure and function occurs during graft remodeling. These investigations have shown as cells migrate into this model ex vivo scaffold significant variation in material elasticity occurs that may have important implications in our understanding of early stage vascular remodeling events.

Introduction

Small diameter vascular bypass graft materials (< 6 mm) have high failure rates due predominantly to poor biological interactions and functionality 1–2. Tissue engineering or de novo regeneration of small diameter blood vessels has shown significant promise to develop biologically functional alternatives as vascular replacements3–4. Significant progress has been made with this approach from the pioneering work of Weinberg and Bell in 1986 using natural hydrogels5 to later examples designed with synthetic materials 6–7 and acellular grafts 8–9. While the potential has been shown with reduced thrombogenicity and enhanced patency rates further progress is needed to provide a routine clinically viable solution 10–11. Of the many candidate materials researched, naturally occurring ex vivo matrices, or polymer derivatives such as collagen or fibronectin, aim to take advantage of native tissue mechanics and biological moieties that enhance biological function and integration 12–16.

An important goal has been to engineer the vascular grafts with similar mechanical properties to natural vessels and as such a significant amount research has been conducted to assess culture systems and environmental conditions that may improve graft performance. In order to recapitulate these conditions, bioreactors and perfusion flow systems aim to emulate the in vivo environment in order to enhance tissue regeneration by modulating cellular phenotype, improving mass-transport limitations, and long-term sterility 17–20.

Using a pulsed perfusion bioreactor, the systems designed by Hoerstrup et al have shown that constructs cultured for 1 month resulted in significant improvements of burst strength, from 50 mmHg for static controls to 326 mmHg for conditioned samples 21. Using a different approach, L’Heureux et al. developed a well defined three-layered vessel structure by adding concentric layers of cellular sheets, that alter twelve weeks of preparation and culture displayed burst strength over 2000 mmHg 22. More recently Syedain et al. investigated the effects of circumferential strain amplitude ranging from 2.5% to 20% either constantly or incrementally to evaluate the effects on fibrin remodeling. It was found that the incremental circumferential strain resulted in higher ultimate tensile strength (2295 ± 467 kPa,50% increase) compared to constant circumferential stress, showing the complex nature of vessel remodeling and adaptation during regenerative processes 23.

While we are beginning to understand the complex nature of tissue remodeling and broad effects of mechanical stimulation, little is currently known of how these early remodeling events influence the biomechanical behavior of these developing tissues. ECM remodeling, and the factors that influence the remodeling process, are clearly important and require more detailed investigation, particularly so with tissue-based materials where the primary ECM macromolecules provide the bulk of the structural support.

Since the late 1970’s the human umbilical vein (HUV) has been used as a glutaraldehyde cross-linked allograft showing improvements in graft patency compared to many synthetic materials24. Similar to other aldehyde treated materials the cross-linking process used on current clinical versions of this material inhibit cell migration and subsequent remodeling due to the inability of cells to degrade these induced bonds. Daniel et al (2005) developed an automated methodology to rapidly and uniformly dissect the HUV directly from umbilical cord25. Using this automated dissection procedure the HUV scaffold used in these investigations was decellularized to form an acellular scaffold without artificially induced stabilization such as formaldehyde or glutaraldehyde treatments. The perceived advantage of this approach a reduced immune response allowing adhered cells (seeded in vitro or naturally in vivo) to positively remodel the scaffold without the inflammatory response rapidly degrading the tissue, which would otherwise lead to graft dilation and possible aneurysm formation. As such remodeling can occur in a more stable fashion leading to a fully cellular, and functional graft.

These investigations used HUV model system to evaluate for blood vessel regeneration. Our experimental design aimed to characterize biomechanical changes that occur during early phase cellular remodeling events with the HUV when prepared as a non-stabilized graft, and compare the variation as a function of dynamic stimulation. To assess these early regenerative events, two independent conditions were evaluated: i) static culture and ii) dynamic perfusion culture. Three sets of perfusion bioreactors were connected in series with independent lumen and ablumenal flow circuits to control perfusion condition. Constructs were cultured to assess changes in construct biomechanics and cellular activity as a function of culture technique, and time.

Materials and Methods

Cell culture

Human myofibroblasts were procured from ATCC (CRL-2854, Virginia, USA) and cultured in high glucose (4.5 g/l) Dulbecco’s Modified Eagle’s Medium (DMEM) (Invitrogen, CA), supplemented with 10% Fetal Bovine Serum (FBS), 1% l-glutamine, and 1% penicillin streptomycin (Invitrogen, CA) with 5% CO2 at 37° C. Cells were prepared for seeding when at 60–70 % confluence at passage 6.

Scaffold preparation and Bioreactor assembly

Veins were dissected from the cord as described previously26, cut to lengths of 110 mm and agitated for 24 h in 1% Triton X-100 on an orbital shaker. Vessels were then rinsed in distilled water for 10 min, followed by 20 min 1 h and 24 h washes. This was followed by incubation in 70 U/ml DNase in PBS supplemented with Mg2+ (Sigma-Aldrich Inc., St. Louis, MO) for 3h at 37°C, then rinsed. HUV segments were then loaded in the perfusion bioreactors (3 in series), as shown in Figure 1, with each scaffold taken from a different placental origin. Three bioreactors (see Figure 1. A–C) were connected in series, with two independent flow circuits (lumen and ablumen), each with respective media reservoirs (see Figure 1. D, E), and rotary pumps (see Figure 1. G, F).

Figure 1. Dynamic cell culture setup.

Figure 1

Open in a new tab

Three bioreactors (A, B, C) were connected in series. Two flow circuits, lumen and ablumen with respective medium reservoirs (D, E) and rotary pumps (G, F). Decellularized HUV samples were seeded with myofibroblasts and cultured for a period of 21 days.

Samples were sterilized using 0.2% peracetic acid and 4% ethanol, circulated through the lumen and albumen sides for 2 h then pH balanced with multiple washes of PBS and pretreated with complete growth media overnight.

Seeding protocol

Cells were seeded to the ablumenal surface of the HUV scaffold using a hydrogel contraction technique developed in our labs. Briefly, human myofibroblasts were suspended in 1.9 mg/ml type 1 collagen gels (Purcol, Inamed, Fremont, CA) containing 1×106 cells/ml and inoculated into the ablumenal void space of the bioreactors surrounding the scaffold (10ml/scaffold). Bioreactors were placed in a 0% CO2 incubator for 5 h at 37°C for gel polymerization. After the initial gel contraction, the culture medium was circulated through the ablumen void space at a flow rate of 1.5 ml/min. After 48 h, lumen flow was slowly ramped in increments of 2.5 ml/min every 2 h until 10 ml/min was reached, at which point ablumen flow was increased to 2 ml/min. Over the following four days the lumenal flow rate was increased in increments of 10 ml/min each day to a maximum of 50 ml/min (90 bpm). Samples cultured under static conditions were seeded as above then placed in T-75 culture flasks and maintained. After 7 and 21 days the vascular constructs were removed from bioreactors and characterized, see Figure 2. A–D.

Figure 2. Hydrogel contraction around the HUV scaffold.

Figure 2

Open in a new tab

The sequence of hydrogel contracting from the bioreactor glass wall and around the HUV is shown. A) The hydrogel was loaded into the bioreactor void space. B) After 5h of gel reaction, the collagen has polymerized and detached from the bioreactor wall. C) The medium was initially perfused around the void space and circulated freely on the ablumen side. D) After the selected time periods, the vascular constructs were removed from bioreactors and characterized.

Construct characterization

Samples were collected each construct (n=3) from the proximal, mid and distal positions (total 9 discrete samples per condition) and were analyzed for cellularity, and cell migration using standard histological techniques, as follows.

Cellular metabolic activity was assayed using the non-toxic redox indicator dye Alamar blue (Biosource International, Camarillo, CA). Alamar blue reagent was added to the sample media (10% dilution) and incubated at 37°C for 6 hours, then analyzed using a Synergy HT plate 15 reader (Bio-Tek, Winooski, VT, USA) to measure the absorbance shift from 570 to 600 nm. DNA concentration was then assessed to determine cell density using the Pico Green assay (Invitrogen, Carlsbad, CA). PicoGreen is a fluorescent nucleic acid stain that binds double stranded DNA (dsDNA) in a stoichiometric ratio, and is used to quantify dsDNA at excitation and emission wavelengths of 485nm and 535nm. Calibration curves were produced for known concentrations of cells to DNA that was then used to determine the DNA concentration/cell. Cellular metabolic activity was normalized against cell density, where the total metabolic activity of each sample was divided into the cell number. Samples for histological analysis were fixed in 3% formaldehyde in PBS, dehydrated, embedded in paraffin blocks, then sectioned at 6 μm. Sections were then stained with H&E, and viewed using a Nikon, Eclipse E800 epifluorescent microscope. Cell migration was measured using Image J (NIH) software, using multiple histology images. The degree of cell migration was assessed from the ablumenal surface to the cells location at each specific time point. Within each section, 9 measurements were taken, and averaged to quantify the average migration for each sample set.

Tensile and cyclic compressive testing

All samples were maintained in complete cell culture media until analysis within 30 minutes of bioreactor disassembly. All mechanical analysis was conducted using a uniaxial tensile testing rig (Instron 5542, MA, USA). For tensile analysis, scaffolds were cut into 5 mm wide ringlets. Tissue specimens (n=9) were loaded using stainless steel L-shaped hooks. Samples were preloaded to a stress of 0.005 N at a rate of 5 mm/min and then elongated until failure, with force and extension recorded over time. To assess the elastic behavior of the vessel wall, material properties were identified over two regions: i) a low strain region which reflects physiological behavior (physiological range), and ii) a high strain region (failure range) where collagen fibers are extended by increasing stress levels until failure 27. The low strain region representing physiological pressure conditions were set to 0.01MPa (80 mmHg) to a maximum of 0.02 MPa (120 mmHg). Material stiffness was calculated from the stress strain data to derive the Young’s Modulus (YM) over these two zones. The slope of the linear region prior to failure was used to calculate the Young’s modulus, measure of material stiffness. The ultimate tensile strength (UTS) was obtained from the stress-strain graph, corresponding to the maximum load that the samples withstood prior to rupture.

Samples prepared for compression analysis were obtained by longitudinally sectioning the vessels then using a circular stainless steel punch removed 0.79 mm OD disks from samples (n=9). Disks were fixed to the lower compression plate and hydrated with PBS. Uniaxial cyclic compression tests were conducted such that the compression vector was perpendicular to the vessel wall. Our investigations assessed the effect of physiologically relevant applied compressive strain (10%) to determine cyclic compressive properties of the constructs28. Indenter displacement was set to 0.02 mm/s and samples compressed to a maximum strain of 10%. The load was then progressively released at the same rate to 0% strain, for a total of 15 cycles. The load versus-displacement data was collected and converted to stress and strain. The maximum value of the compressive stiffness was derived from the maximum slope of the stress-versus-strain response (up to 10% strain) during compression loading. Energy dissipation per unit volume was calculated from the hysteresis data for each construct. Mean values of energy dissipation per unit volume in the final compression cycle were compared to the initial cycle, and calculated as a percentage.

Statistical analysis

All analyses were carried out in triplicate unless otherwise stated. From each of the 3 constructs comprising the triplicate set, a further 3 samples were analyzed, obtaining a total of nine data points for each condition. A multi comparison Tukey test by one-way ANOVA (p <0.05) was used to determine statistical significance. All data are presented as the mean ± standard deviation.

Results

Construct thickness

Construct thickness was measured in proximal, mid and distal position and the average is presented in Figure 3. Constructs exposed to dynamic culture conditions for 7 and 21 days displayed a decrease in wall thickness relative to both static culture constructs and acellular static controls. The wall thickness of constructs cultured under static conditions and acellular controls did not show any significant change. The initial average value for the wall thickness was 1.8 mm. Dynamic conditioned constructs at day 7 had an average wall thickness of 1.08 ± 0.04 mm, whereas static cultures yielded 1.72 ± 0.3 mm. At day 21, wall thickness was 1.28 ± 0.15 mm and 1.6 ± 0.19 mm for dynamic and static respectively.

Figure 3. Constructs thickness.

Figure 3

Open in a new tab

The initial average value for the wall thickness is represented with the dashed line. Dynamic cultured constructs exhibited reduced thickness from the static samples, as well as from the initial thickness value.

Cell density

After 7 days, no significant difference was detected in cellularity between dynamic and static constructs (p value= 0.579). However in response to longer periods of dynamic stimulation (21 days), cell growth increased significantly from 0.99×106 ± 0.18 cells/10 mm ringlet to 1.94×106 ± 0.47 cells/10 mm, see Figure 4. A (p value=0.003).

Figure 4.

Figure 4

Open in a new tab

Cell density (A), and metabolic activity (B) of constructs cultured under dynamic and static conditions over 7 and 21 days.

Cellular metabolic activity

AB reduction was assessed to evaluate the metabolic activity of cells adhered to the scaffold. Data for gross metabolic activity was combined with quantitative cell density and expressed as the percentage of AB reduction per cell and shown in Figure 4. B. HUV constructs cultured under dynamic stimulation conditions within the bioreactors displayed a significant increase in metabolic activity over static samples at both 7 and 21 days. However, the culture time had no significant effect on activity, where similar values were found for both culture methods at day 21.

Histological analysis

Cell migration and location within the HUV constructs displayed in Figure 5, showing that distribution within the HUV was strongly dependant on the culture approach. After 7 days of dynamic culture, cells formed continuous interconnected layers cells, around the periphery of the scaffold with infiltration up to 44±11 μm. By contrast, after 21 days of perfusion flow conditioning, cells displayed increased cellular in-growth and migrated into the scaffold up to 101±20 μm, forming clusters of increased cell density embedded within the matrix pores. The overall structure of the HUV was largely retained showing a defined circularly disposed tunica media, followed by a layer with less packed collagen fibers or adventitia leading to an outer layer corresponding to the Wharton’s Jelly material from the cord. Samples exposed to dynamic culture had a reduced wall thickness, displaying a more compressed structure, see Figure 3 and 5.

Figure 5. Histological images of statically and dynamically conditioned HUV constructs.

Figure 5

Open in a new tab

Cells cultured under static conditions where shown to be more disperse and at a lower density relative to dynamic cultures, with cells penetrating up to 200 μm. Dynamic conditioning resulted in an increase in cell concentration, both in unit area and overall total density.

Tensile analysis

No statistical difference in UTS was noted between the two culture conditions at day 21 days (p=0.703); however, values were lower than acellular controls at 450 kPa. The construct UTS at day 7 was shown to be 220 ± 60 kPa and 140 ± 20 kPa, and at day 21 to be 180 ± 60 kPa, and 180 ± 40 kPa, for static and dynamic cultures respectively, see Figure 6.

Figure 6. Ultimate tensile strength (UTS).

Figure 6

Open in a new tab

Mechanical properties of constructs were evaluated at 7 and 21 days in culture. No significant difference in the ultimate tensile strength was found between statically and dynamically conditioned vascular constructs (p>0.05) using one-way ANOVA. Dashed line denotes acellular HUV control.

Mechanical stiffness (tissue elasticity) in the low strain region of the dynamically stimulated constructs increased from 450 ± 140 kPa to 550 ± 190 kPa over the culture period. By 21 days both static and dynamic scaffolds had statistically similar values showing an increase in stiffness relative to acellular controls. By contrast the elasticity of dynamically conditioned constructs in the failure range increased 66% from 610 ± 210 kPa at day 7 to 200 ± 110 kPa at day 21. The YM of static cultures remained constant at 550 ± 60 and 790 ± 280 kPa for days 7 and 21 respectively, see Figure 7.

Figure 7. Comparison of Young’s modulus, and tensile strain at maximum stress over the physiological range (left), and failure range (right).

Figure 7

Open in a new tab

Dynamic culture resulted in more elastic constructs after day 21, whereas under static culture, this property remained unchanged. After 21 days, the dynamic culture group presented decreased stiffness compared to static group (p>0.05). Dashed line denotes acellular HUV control..

Construct elongation, expressed as the tensile strain (%) and tensile strain at max load was calculated at i) physiological range and, ii) failure range. Figure 7 shows the physiological range with a max stress value of 120 mmHg (0.02 MPa), and the failure range prior to failure. Over the physiological range, the statically cultured constructs displayed increased strain values compared to dynamic constructs. In the failure range, results show constructs exposed to dynamic conditioning have a significant increase in strain value from day 7 (82 ± 20.5%) to day 21 (227 ± 59.5%). Static constructs remained unchanged.

Compression analysis

A representative stress-strain response curve for samples under compressive loading is represented in Figure 8. A. The response of the material to cyclic compression displayed the characteristic soft tissue hysteresis associated with energy dissipation as the cyclic stress is released. For both groups, dynamic and static, the tissue resistance to compression and energy dissipation was the highest during the first compressive cycle and then it decreased in subsequent cycles. The mean values of peak (kj/m3) hysteresis in the final compression cycle was compared to the initial cycle. Data shows energy dissipation after the first cycle and in subsequent cycles to reduce up to 90% of the original value by the 15th cycle, see Figure 8. B. Only dynamically stimulated constructs at day 21 showed a statistically significant reduction compared to the other constructs.

Figure 8.

Figure 8

Open in a new tab

(A) Representative compressive stress–strain relationship for the HUV constructs under cyclic compression. Dynamic samples displayed resistance to compression from an applied strain of 5%. The hysteresis loops indicate that energy is dissipated inside the disk when load is retrieved. (B) % energy dissipation. The mean values of hysteresis in the final compression cycle as compared to the initial cycle were plotted. The amount of energy dissipation in subsequent cycles was reduced to about 50% to 90% of the original value after 15 cycles. Dashed line denotes acellular HUV control.

The maximum compressive strength reached during the first cycle at day 7 was not significantly different than at day 21 for both culture conditions. Similarly, the response to the last compressive cycle was unchanged over the evaluated time period, see Figure 9. The compressive stiffness was calculated from the stress strain curves as the maximum slope of the compressive stress-strain graph in the first cycle remained unchanged from day 7 to day 21 under both dynamic and static conditions, see Figure 9. At day 21 constructs had a compressive modulus of 20.7 ± 7.0 kPa under static conditions, and 23.3 ± 8.0 kPa for dynamic conditions.

Figure 9. Maximum strength obtained during first and last compressive cycles.

Figure 9

Open in a new tab

At day 21, significant difference was presented between the peak strength at first and last cycles. Dashed line denotes acellular HUV control. Compressive stiffness of the vascular constructs. The mean compressive was obtained as the ratio of the strength over strain at the maximum strain value. No significant difference was obtained between groups. Dashed line denotes acellular HUV control.

Discussion

A critical, yet poorly understood, aspect of the remodeling processes is the biomechanical transformation that occurs throughout regenerative process. It has been shown that when material mechanical properties are miss-matched to the host vasculature (at the implantation site) it is predictive of graft failure, and as such is a critical parameter to assess 29–30. From this perspective ex vivo-based materials have an advantage over currently used synthetic conduits that are typically stiffer and less compliant than native vessels. With ex vivo materials the body’s natural remodeling machinery progressively modifies the ECM through balanced degradative and synthetic pathways that alter the tissues mechanical properties over time. A balance between ECM synthesis and degradation is clearly a critical factor, particularly with arterial grafts where ECM failure may have fatal consequences.

In these investigations, human myofibroblasts were seeded onto an ex vivo-based acellular scaffold and cultured under dynamic perfusion conditions to assess broad biological phenomena in concert with biomechanical changes. By assessing these properties our goal was to to further our understanding of how in vitro culture influences graft mechanics during early phase remodeling events. A number of in-vitro studies by other groups have demonstrated the effects of mechanical stimulation and the direct effects on accelerated cell proliferation, extracellular matrix production and vascular remodeling 31–33. Our investigations also confirm this broad trend where constructs cultured under dynamic conditions were shown to have higher metabolic activity and cellularity than comparative constructs cultured under static conditions. Similarly, Stegemann and Nerem31 have shown that embedding SMCs in 3D collagen scaffolds, without mechanical loading, leads to decreased cellular proliferation and α-actin expression relative to monolayer cultures. When subjected to a low load environment (i.e. floating collagen matrix) cells were inclined to become quiescent over time and as the mechanical load was increased, fibroblasts displayed higher proliferation rates, matrix synthesis and α-actin fiber formation. Mechanical stimulation has generally been shown to benefit construct properties. However in vitro, extended culture times may not result in beneficial remodeling. The investigations conducted by Seliktar et al. showed that while mechanically stimulated collagen constructs seeded with human aortic SMCs displayed enhanced ultimate tensile stress and increased stiffness values over shorter culture periods (4 days), constructs subjected to prolonged mechanical stimulation (8 days) deteriorated. Based on their findings, it was suggested that the variation in collagen compaction and consequent mechanical properties is the result of a cell-mediated remodeling that is largely regulated by mechanical stimulation 34. While these time points are reduced compared to our study there is some correlation with a change in stiffness that maybe attributed to a similar mechanism where the balance between remodeling degradation and synthesis may not always be in balance. This again demonstrates the importance of mechanical and biochemical cues that determine cell phenotype 31.

A materials compression characteristics is a physical property often overlooked that provides a better understanding of a materials bulk properties - over and above what tensile analysis offers. Broadly, mechanical analysis can be put into two data sets. The first is to compare a materials behavior (in this case vascular tissue) to relevant physiological conditions, and the secondly to understand how a material performs, as a discrete material, irrespective of its application. Tensile testing can represent ‘stretching, or stress loading’ of a tissue, it also characterizes performance that allows others to compare properties to other materials. This is also the case with compression analysis. Compressive stress is an important parameter of wall mechanics over the systolic/diastolic pressure change, as indicated by a thinning of the wall as pressure increases35. Again, while this is a useful physiological attribute to assess, it plays a significant role in a more fundamental characterization of the materials properties that allows direct comparison.

As part of this compressive testing the energy dissipation was calculated to understand how these vascular grafts soften during cyclic loading. As a function of how fluid is squeezed out during compression, and how it returns as pressure is released relative to the initial cyclic load. A slower recovery (toward its initial condition) infers the tissue has become softer, relative to a control sample36. In this study, for dynamically stimulated constructs at day 21, it is interesting that while the scaffolds elasticity increased at failure range, and % reduction of energy dissipation showed significant reduction, ‘therefore softening of these constructs’. No change was noted in the UTS and in compressive stiffness. This variation was only noted in the failure range with the YM in the physiological range remaining unchanged at day 21 with dynamically cultured constructs. Increased stiffness at low strain region may be attributed to partial fragmentation of elastin fibers that results with the increased contribution of collagen fibers37. At the higher strain values (failure range) a number of possibilities may explain the 66 % decrease in stiffness but without any significant change in the UTS. A partial degradation of the bonds that hold the larger collagen fibers together, but not of the actual fibrils themselves would largely maintain the ECM strength but may decrease stiffness as molecules are freer to slide against one another. This may be further attenuated by the outer Wharton’s jelly that consists largely of GAG’s is remodeled with a higher (relative) concentration of collagen. This may maintain the UTS, but as the collagen has not fully matured (low cross-link density) increased stiffness results.

In vivo, graft dilatation may occur as early as the initial adaptation period where overexpression of MMP’s by vascular smooth muscle cells and inflammatory cells can result in degradation of elastin and collagen38. To some extent weakening of the scaffold is expected during this early remodeling period as the newly synthesized and deposited ECM is in a less structured orientation. It is then reorganized as the remodeling process matures and naturally cross-linked to enhance the strength of the collagen fibers and ECM as a whole. However a secondary mechanism may explain a partial ECM deterioration that is more specific to acellular (ex vivo) scaffolds. It is hypothesized that expressed MMP’s that are typically associated closely with the cell membrane may have a more distant effect of ECM mechanics by freely diffusing through the acellular scaffold to more distant zones of the scaffold. While there is no direct evidence for this effect, this may explain an overall weakening that may occur, especially if cell migration is limited. Further investigation is required to detail these possibilities.

Engineering vessels that possess similar biomechanical properties to natural arteries is an important research objective. Scaffold remodeling, the key process in effective regeneration, is a complex process that is clearly influenced by multiple factors including mechanical stimulation, scaffold composition and structure, cell type, culture chemistry as well as many other more subtle elements. Mismatch between graft and the host artery causes a variety of effects that can result in abnormal endothelial and smooth muscle cell phenotype that results in pathologies such as thrombosis and intimal hyperplasia, which are primary causes of graft failure 39–40. In this study mechanical conditioning is used to direct cell function, and as a result biomechanical properties of the tissue scaffold changed overtime. Maintaining, or at least directing the biomechanical changes that occur during scaffold remodeling remain a challenge. These findings have shown that studying the HUV biomechanical properties of engineered blood vessels under conditions that broadly mimic in vivo mechanics is a useful model to better understand in vitro remodeling processes. While there are some parallels that can be drawn with in vivo remodeling, these in vitro investigations are designed to be more progressive in nature where single cell populations can be assessed independently. As such, the remodeling processes in vivo are considerably more complex with contributions by other cellular systems, including the host immune response. Further optimization will provide insight into these more complex remodeling processes that may improve our understanding of both heterologous transplant graft remodeling and engineered vessel regeneration.

Acknowledgments

We gratefully acknowledge financial support of the National Institutes of Health (NIH) for their support – grant: R01HL0882

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  • 1.Wang X, Lin P, Yao Q, Chen C. Development of small-diameter vascular grafts. World J Surg. 2007;31(4):682–9. doi: 10.1007/s00268-006-0731-z. [DOI] [PubMed] [Google Scholar]
  • 2.Kakisis JD, Liapis CD, Breuer C, Sumpio BE. Artificial blood vessel: the Holy Grail of peripheral vascular surgery. J Vasc Surg. 2005;41(2):349–54. doi: 10.1016/j.jvs.2004.12.026. [DOI] [PubMed] [Google Scholar]
  • 3.Zhang WJ, Liu W, Cui L, Cao YL. Tissue engineering of blood vessel. Journal of Cellular and Molecular Medicine. 2007;11(5):945–57. doi: 10.1111/j.1582-4934.2007.00099.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Thomas AC, Campbell GR, Campbell JH. Advances in vascular tissue engineering. Cardiovasc Pathol. 2003;12(5):271–6. doi: 10.1016/s1054-8807(03)00086-3. [DOI] [PubMed] [Google Scholar]
  • 5.Weinberg CB, Bell E. A blood vessel model constructed from collagen and cultured vascular cells. Science. 1986;231(4736):397–400. doi: 10.1126/science.2934816. [DOI] [PubMed] [Google Scholar]
  • 6.Matsuda T, Akutsu T, Kira K, Matsumoto H. Development of hybrid compliant graft: rapid preparative method for reconstruction of a vascular wall. ASAIO Trans. 1989;35(3):553–5. [PubMed] [Google Scholar]
  • 7.Kottke-Marchant K, Anderson JM, Umemura Y, Marchant RE. Effect of albumin coating on the in vitro blood compatibility of Dacron arterial prostheses. Biomaterials. 1989;10(3):147–55. doi: 10.1016/0142-9612(89)90017-3. [DOI] [PubMed] [Google Scholar]
  • 8.Yu XX, Wan CX, Chen HQ. Preparation and endothelialization of decellularised vascular scaffold for tissue-engineered blood vessel. J Mater Sci Mater Med. 2008;19(1):319–26. doi: 10.1007/s10856-007-3157-8. [DOI] [PubMed] [Google Scholar]
  • 9.Daniel J, Abe K, McFetridge PS. Development of the human umbilical vein scaffold for cardiovascular tissue engineering applications. ASAIO J. 2005;51(3):252–61. doi: 10.1097/01.mat.0000160872.41871.7e. [DOI] [PubMed] [Google Scholar]
  • 10.McGuigan AP, Sefton MV. The thrombogenicity of human umbilical vein endothelial cell seeded collagen modules. Biomaterials. 2008;29(16):2453–63. doi: 10.1016/j.biomaterials.2008.02.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Seifalian AM, Tiwari A, Hamilton G, Salacinski HJ. Improving the clinical patency of prosthetic vascular and coronary bypass grafts: the role of seeding and tissue engineering. Artif Organs. 2002;26(4):307–20. doi: 10.1046/j.1525-1594.2002.06841.x. [DOI] [PubMed] [Google Scholar]
  • 12.McFetridge PS, Abe K, Horrocks M, Chaudhuri JB. Vascular tissue engineering: Bioreactor design considerations for extended culture of primary human vascular smooth muscle cells. Asaio Journal. 2007;53(5):623–30. doi: 10.1097/MAT.0b013e31812f3b7e. [DOI] [PubMed] [Google Scholar]
  • 13.Borschel GH, Huang YC, Calve S, Arruda EM, Lynch JB, Dow DE, et al. Tissue engineering of recellularized small-diameter vascular grafts. Tissue Eng. 2005;11(5–6):778–86. doi: 10.1089/ten.2005.11.778. [DOI] [PubMed] [Google Scholar]
  • 14.Lalka SG, Oelker LM, Malone JM, Duhamel RC, Kevorkian MA, Raper BA, et al. Acellular vascular matrix: a natural endothelial cell substrate. Ann Vasc Surg. 1989;3(2):108–17. doi: 10.1016/S0890-5096(06)62002-5. [DOI] [PubMed] [Google Scholar]
  • 15.Malone JM, Brendel K, Duhamel RC, Reinert RL. Detergent-extracted small-diameter vascular prostheses. J Vasc Surg. 1984;1(1):181–91. doi: 10.1067/mva.1984.avs0010181. [DOI] [PubMed] [Google Scholar]
  • 16.Wilson GJ, Courtman DW, Klement P, Lee JM, Yeger H. Acellular matrix: a biomaterials approach for coronary artery bypass and heart valve replacement. Ann Thorac Surg. 1995;60(2 Suppl):S353–8. doi: 10.1016/0003-4975(95)98967-y. [DOI] [PubMed] [Google Scholar]
  • 17.McFetridge PS, Bodamyali T, Horrocks M, Chaudhuri JB. Endothelial and smooth muscle cell seeding onto processed ex vivo arterial scaffolds using 3D vascular bioreactors. ASAIO J. 2004;50(6):591–600. doi: 10.1097/01.mat.0000144365.22025.9b. [DOI] [PubMed] [Google Scholar]
  • 18.Bjork JW, Tranquillo RT. Transmural flow bioreactor for vascular tissue engineering. Biotechnology and Bioengineering. 2009;104(6):1197–206. doi: 10.1002/bit.22475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Chen HC, Hu YC. Bioreactors for tissue engineering. Biotechnol Lett. 2006;28(18):1415–23. doi: 10.1007/s10529-006-9111-x. [DOI] [PubMed] [Google Scholar]
  • 20.Villalona GA, Udelsman B, Duncan DR, McGillicuddy E, Sawh-Martinez RF, Hibino N, et al. Cell-Seeding Techniques in Vascular Tissue Engineering. Tissue Engineering Part B: Reviews. 2010;16(3):341–50. doi: 10.1089/ten.teb.2009.0527. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Hoerstrup SP, Zund G, Sodian R, Schnell AM, Grunenfelder J, Turina MI. Tissue engineering of small caliber vascular grafts. Eur J Cardiothorac Surg. 2001;20(1):164–9. doi: 10.1016/s1010-7940(01)00706-0. [DOI] [PubMed] [Google Scholar]
  • 22.L’Heureux N, Paquet S, Labbe R, Germain L, Auger FA. A completely biological tissue-engineered human blood vessel. FASEB J. 1998;12(1):47–56. doi: 10.1096/fasebj.12.1.47. [DOI] [PubMed] [Google Scholar]
  • 23.Syedain ZH, Weinberg JS, Tranquillo RT. Cyclic distension of fibrin-based tissue constructs: evidence of adaptation during growth of engineered connective tissue. Proc Natl Acad Sci U S A. 2008;105(18):6537–42. doi: 10.1073/pnas.0711217105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Dardik H. The second decade of experience with the umbilical vein graft for lower-limb revascularization. Cardiovascular Surgery. 1995;3(3):265–9. doi: 10.1016/0967-2109(95)93874-o. [DOI] [PubMed] [Google Scholar]
  • 25.Daniel J, Abe K, McFetridge PS. Development of the Human Umbilical Vein Scaffold for Cardiovascular Tissue Engineering Applications. ASAIO Journal. 2005;51(3):252–61. doi: 10.1097/01.mat.0000160872.41871.7e. [DOI] [PubMed] [Google Scholar]
  • 26.Montoya CV, McFetridge PS. Preparation of Ex Vivo–Based Biomaterials Using Convective Flow Decellularization. Tissue Engineering Part C: Methods. 2009;15(2):191–200. doi: 10.1089/ten.tec.2008.0372. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Duprey A, Khanafer K, Schlicht M, Avril S, Williams D, Berguer R. In vitro characterisation of physiological and maximum elastic modulus of ascending thoracic aortic aneurysms using uniaxial tensile testing. Eur J Vasc Endovasc Surg. 2010;39(6):700–7. doi: 10.1016/j.ejvs.2010.02.015. [DOI] [PubMed] [Google Scholar]
  • 28.Mauck RL, Soltz MA, Wang CCB, Wong DD, Chao P-HG, Valhmu WB, et al. Functional Tissue Engineering of Articular Cartilage Through Dynamic Loading of Chondrocyte-Seeded Agarose Gels. Journal of Biomechanical Engineering. 2000;122(3):252–60. doi: 10.1115/1.429656. [DOI] [PubMed] [Google Scholar]
  • 29.Sarkar S, Salacinski HJ, Hamilton G, Seifalian AM. The mechanical properties of infrainguinal vascular bypass grafts: their role in influencing patency. Eur J Vasc Endovasc Surg. 2006;31(6):627–36. doi: 10.1016/j.ejvs.2006.01.006. [DOI] [PubMed] [Google Scholar]
  • 30.Seifalian AMAG, Schmitz-Rixen T, Hamilto G. Noncompliance: The Silent Acceptance of a Villain. In: Zilla P, editor. Tissue Engineering of Vascular Prosthetic Grafts. Landes Bioscience; 1991. [Google Scholar]
  • 31.Stegemann JP, Nerem RM. Phenotype Modulation in Vascular Tissue Engineering Using Biochemical and Mechanical Stimulation. Annals of Biomedical Engineering. 2003;31(4):391–402. doi: 10.1114/1.1558031. [DOI] [PubMed] [Google Scholar]
  • 32.Niklason LE, Gao J, Abbott WM, Hirschi KK, Houser S, Marini R, et al. Functional arteries grown in vitro. Science. 1999;284(5413):489–93. doi: 10.1126/science.284.5413.489. [DOI] [PubMed] [Google Scholar]
  • 33.Xu ZC, Zhang WJ, Li H, Cui L, Cen L, Zhou GD, et al. Engineering of an elastic large muscular vessel wall with pulsatile stimulation in bioreactor. Biomaterials. 2008;29(10):1464–72. doi: 10.1016/j.biomaterials.2007.11.037. [DOI] [PubMed] [Google Scholar]
  • 34.Seliktar D, Nerem RM, Galis ZS. Mechanical strain-stimulated remodeling of tissue-engineered blood vessel constructs. Tissue Eng. 2003;9(4):657–66. doi: 10.1089/107632703768247359. [DOI] [PubMed] [Google Scholar]
  • 35.Chaudhuri J, Al-Rubeai M. Bioreactors for tissue engineering: principles, design and operation. Dordrecht: Springer; 2005. [Google Scholar]
  • 36.Lumpkins SB, McFetridge PS. Regional variations in the viscoelastic compressive properties of the temporomandibular joint disc and implications toward tissue engineering. J Biomed Mater Res A. 2009;90(3):784–91. doi: 10.1002/jbm.a.32148. [DOI] [PubMed] [Google Scholar]
  • 37.Fonck E, Prod’hom G, Roy S, Augsburger L, Rüfenacht DA, Stergiopulos N. Effect of elastin degradation on carotid wall mechanics as assessed by a constituent-based biomechanical model. American Journal of Physiology - Heart and Circulatory Physiology. 2007;292(6):H2754–H63. doi: 10.1152/ajpheart.01108.2006. [DOI] [PubMed] [Google Scholar]
  • 38.Busti C, Falcinelli E, Momi S, Gresele P. Matrix metalloproteinases and peripheral arterial disease. Internal and Emergency Medicine. 2010;5(1):13–25. doi: 10.1007/s11739-009-0283-y. [DOI] [PubMed] [Google Scholar]
  • 39.Chan-Park MB, Shen JY, Cao Y, Xiong Y, Liu Y, Rayatpisheh S, et al. Biomimetic control of vascular smooth muscle cell morphology and phenotype for functional tissue-engineered small-diameter blood vessels. Journal of Biomedical Materials Research Part A. 2009;88A(4):1104–21. doi: 10.1002/jbm.a.32318. [DOI] [PubMed] [Google Scholar]
  • 40.Salacinski HJ, Goldner S, Giudiceandrea A, Hamilton G, Seifalian AM, Edwards A, et al. The Mechanical Behavior of Vascular Grafts: A Review. J Biomater Appl. 2001;15(3):241–78. doi: 10.1106/NA5T-J57A-JTDD-FD04. [DOI] [PubMed] [Google Scholar]

RESOURCES