Introduction

The lack of organs and tissues is still a ubiquitous challenge of international healthcare today. To overcome this problem, tissue engineering has become a major field of medical research in recent years. Since only a sufficient blood supply can meet the metabolic demands of living tissues, the immediate development of a dense and functional microvascular network is crucial for the successful incorporation of tissue engineering constructs [1].

Different strategies have evolved to accelerate blood vessel ingrowth into tissue engineering constructs. Direct single applications as well as combinations of angiogenic growth factors have been applied [2,3,4]. Attempts to generate or prefabricate vascular structures within tissue engineering constructs have also been performed [5,6,7]. However, the vitalization of suitable scaffolds with pluripotent (e.g., bone marrow mesenchymal stem cells, bmMSCs) or organ-specific cells (e.g., osteoblast-like cells, OLCs) remains the basic principle [8, 9]. Several of these approaches have shown promising results, but two fundamental problems hamper a successful transfer into the clinic. First, most of the studies have been conducted in vitro. Second, even though the quality of the local host tissue plays a key role in vascularization, healthy conditions have been chosen in almost all previous projects [10]. However, except for patients with traumatic destruction or rare diseases, tissue engineering constructs are mainly required for old and multimorbid patients. These patients often suffer from diseases that limit the capacity of neovascularization, e.g., diabetes or hypertension [11]. Arterial hypertension is the most common internal disease worldwide [12]. At least 15 million people die annually from stroke and ischemic heart disease. In this regard, it is assumed that approximately 50% of strokes and myocardial infarctions are caused by hypertension [13, 14]. At this time, 25% of the global population suffers from elevated blood pressure. By 2025, this number will have grown to 29% (1.5 billion people) [15]. Referring to the “Global Burden of Disease Study” from 2015, hypertension is the most important risk factor concerning quality and expectation of life [16].

Regarding microcirculation in the hypertensive organism, various aspects have to be considered. Among other adjustment processes to high blood pressure, small arteries and arterioles react with eutrophic inward remodeling, in which the vessel wall components are rearranged without growth [17, 18]. Microvascular rarefaction has been observed over many years in hypertensive patients and animal models [17, 19, 20]. Moreover, capillary recruitment is significantly reduced in hypertensive patients, and recruitment is inversely correlated with increasing blood pressure [17, 21]. Additionally, blood rheology is hindered due to the high incidence of circulating leukocytes under hypertensive conditions [17, 22].

Therefore, we investigated the in vivo vascularization capacity of tissue engineering constructs in a hypertensive organism. For this purpose, poly-L-lactide-co-glycolide (PLGA) scaffolds were seeded with organ-specific (OLCs) or undifferentiated progenitor cells (bmMSCs) and implanted into the dorsal skinfold chambers of hypertensive BPH/2J mice. Normotensive BPN/3J mice served as controls. After the transplantation of the tissue engineering constructs, we analyzed the angiogenic and inflammatory parameters using repetitive intravital fluorescence microscopy over a time period of 2 weeks.

Methods

The current study represents a further development of a large-scale and continuous study performed by the authors on the early acceleration of the vascularization of tissue engineering constructs in vivo [2, 3, 23,24,25,26,27,28,29]. To guarantee the comparability of results within this experimental series, the methodology was similar to that of our previous studies. Therefore, the materials and methods are briefly summarized with references to further details.

Animals

The experiments were conducted in accordance with the German legislation for the protection of animals and the NIH Guidelines for the Care and Use of Laboratory Animals (NIH Publication No. 85-23 Rev. 1985). The experiments were approved by the local governmental animal care committee (Niedersächsisches Landesamt für Verbraucherschutz und Lebensmittelsicherheit, reference number 33.9-42502-04-08/1437). Male hypertensive BPH/2J mice were used for the experiments at the age of approximately 20 weeks and a body weight of 16–27 g (Jackson Laboratories, Bar Harbor, USA). An increased systolic blood pressure (134 ± 8 mm Hg), an accelerated heart rate and a heightened hematocrit characterize this inbred mouse strain [30, 31]. Male BPN/3J mice (Jackson Laboratories) with a systolic blood pressure of 92 ± 6 mm Hg were used as controls (Table 1). BPN/3J represents the nonselective reproduced control strain to BPH/2J. The keeping of animals was conducted according to a standard protocol [23,24,25].

Table 1 Basic characteristics of mice
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Preparation of the dorsal skinfold chamber

Surgery, scaffold implantation and repetitive intravital fluorescence microscopy were performed under well-established anesthetic procedures [23,24,25].

The dorsal skinfold chamber allowed intravital microscopic surveillance of microcirculation in the anesthetized animal over an extended period. The surgical techniques for the preparation of the dorsal skinfold chamber and scaffold implantation have been previously described [32]. Within this experimental setup, PLGA scaffolds were implanted after a minimum 48-h recovery period subsequent to surgical preparation.

Fabrication of tissue engineering constructs

PLGA scaffolds with dimensions of 3 × 3 × 1 mm were fabricated at the Freiburg Institute for Material Research and Macromolecular Chemistry as previously described [33].

To seed PLGA scaffolds with OLCs, cells were isolated from male BPN/3J mice, age 8 weeks, as presented recently [23]. OLCs were characterized by rabbit anti-mouse collagen I (Biotrend, Köln, Germany), rabbit anti-mouse osteocalcin (Acris Antibodies GmbH, Hiddenhausen, Germany) and rat anti-mouse osteonectin (SPARC, R&D Systems, Wiesbaden, Germany) [34]. For additional characterization of OLCs, histochemical alkaline phosphatase determination was performed [35].

Correspondingly, bone marrow was obtained from male BPN/3J mice, age 8 weeks, and bone-marrow-derived murine mesenchymal stem cells were isolated according to established protocols based on plastic adherence of these cells [23, 36, 37]. Mesenchymal stem cells were characterized with anti-mouse Ly-6A/E (Sca-1) (Biolegend, San Diego, USA) and anti-mouse CD44 (Biolegend) [37]. Cy2-conjugated goat anti-rabbit antibody and Cy2-conjugated goat anti-rat antibody (both Dianova, Hamburg, Germany) were used as secondary antibodies. Nuclei were stained using DAPI (Sigma, Darmstadt, Germany).

For in vivo experiments, PLGA scaffolds were coated with collagen type I (Biotrend, Köln, Germany) according to standardized guidelines [23]. Cultivated cells were rinsed three times with HBSS and then incubated with Accutase (OLCs) or Trypsin/EDTA (bmMSCs) (PAA, Coelbe, Germany) until cells were released from the culture surface. Cells were washed twice with culture medium and then counted. Subsequently, 1 × 104 OLCs or bmMSCs were resuspended in 10 μL of culture medium and spotted onto the PLGA scaffolds coated with collagen type I. The scaffolds with cells were incubated at 37 °C in a humidified incubator in an atmosphere with 91.5% air and 8.5% CO2 for 4 h to ensure cell attachment to the scaffold surfaces. Scaffolds with attached cells were placed in fresh culture medium and incubated overnight at 37 °C in a humidified incubator in an atmosphere with 95% air and 5% CO2. Afterwards, the prepared scaffolds were implanted into the dorsal skinfold chambers of BPH/2J and BPN/3J mice.

Intravital fluorescence microscopy

For intravital fluorescence microscopy, mice were anesthetized and immobilized with a custom-made stereotactic frame (Central Research Laboratory, Hannover Medical School). After an intravenous injection of 0.1 mL of 5% fluorescein isothiocyanate (FITC)-labeled dextran 150,000 (Sigma) for contrast enhancement by the staining of blood plasma and 0.1 mL of 0.1% rhodamine 6G (Sigma) for direct staining of white blood cells, intravital fluorescence microscopy was performed using a modified Zeiss Axiotech microscope equipped with a 100 W mercury lamp attached to an illuminator with blue, green and ultraviolet filter blocks (Zeiss, Jena, Germany) for epi-illumination. The microscopic images were recorded by a charge-coupled device video camera (FK-6990, COHU, Prospective Measurements, San Diego, USA) and transferred to a DVD recorder (LQ-MS 800, Panasonic, Hamburg, Germany) for off-line evaluation. With the use of 10x and 20x long-distance objectives, the images were displayed on a 14 inch video screen (Panasonic, DVD video recorder LQ-MD 800E, Matsushita Electronic Industrial Co. Ltd., Osaka, Japan).

Microcirculatory analysis

Quantitative off-line analysis of the DVDs was performed by means of the computer-assisted image analysis system CapImage (Zeintl, Heidelberg, Germany). Leukocyte-endothelial cell interaction, microhemodynamics and macromolecular leakage were assessed at a magnification of 20x in 4 different microvascular regions of interest (ROIs) in the border zone of the scaffolds. In each ROI, 1-3 venules (inner diameter: 20-40 µm) were selected for measurements. Leukocytes were classified according to their interaction with the vascular endothelium as adherent, rolling or free-flowing cells as described previously [38]. Adherent leukocytes were defined in each vessel segment as cells that did not move or detach from the endothelial lining within a specified observation period of 20 s and are given as the number of cells per square millimeter of endothelial surface, calculated from the diameter and length of the vessel segment studied, assuming a cylindrical vessel geometry [39]. Rolling leukocytes were defined as cells moving with a velocity less than two-fifths of the centerline velocity and are given as the number of cells per minute, passing a reference point within the microvessel. Diameters, centerline velocity, volumetric blood flow, and wall shear rate were determined in those venules in which leukocyte-endothelial cell interaction was analyzed. Diameters (d) were measured in μm perpendicular to the vessel path. Centerline red blood cell velocity (v) was analyzed by the computer-assisted image analysis system using the line shift method. Volumetric blood flow was calculated by Q = π x (d/2)2 × v/1.6 (pl/s), where 1.6 represents the Baker-Wayland factor to correct for the parabolic velocity profile in microvessels with diameters >20 μm [40]. The wall shear rate (y) was calculated based on the Newtonian definition: y = 8 × v/d. Macromolecular leakage served as an indicator for microvascular permeability and was assessed after an intravenous injection of the macromolecular fluorescent dye FITC-labeled dextran 150,000 by densitometrically determining the gray levels in the tissue directly adjacent to the venular vessel wall (E1) and in the marginal cell-free plasma layer within the vessel (E2). Extravasation (E) was then calculated as E = E1/E2.

Angiogenesis was assessed at 200x magnification. For this purpose, functional microvessel density was measured quantitatively in 8 regions of interest (ROIs) in the border zone of the scaffolds and in 4 regions of interest (ROIs) in the center of the scaffolds. Microvessels that showed a distinct ingrowth, i.e., new capillary buds and sprouts, were counted. By computer-assisted analysis using CapImage software, the cumulative length of microvessels per observation area given in cm/cm2 was calculated [41]. For microcirculatory analysis, investigators were blinded to the treatment group.

In vivo animal experiments

For intravital fluorescence microscopy, 24 BPH/2J mice were randomly divided into three groups with eight animals in each group. Each group was equipped with dorsal skinfold chambers. The PLGA scaffolds were coated with collagen (n = 8), coated with collagen and seeded with OLCs (n = 8), or coated with collagen and seeded with bmMSCs (n = 8) and then implanted. BPN/3J mice, also randomly divided into three groups with eight animals in each group, served as controls. During implantation, care was taken to avoid contamination, mechanical disruption, and chamber damage. The macroscopic appearance of the skinfold chamber preparations and the implanted tissue engineering constructs was documented daily. Intravital fluorescence microscopy of the microhemodynamics, leukocyte-endothelial cell interaction, macromolecular leakage, and angiogenesis was performed immediately and 3, 6, 10, and 14 days after the implantation.

At the end of the in vivo experiments, the animals were euthanized with an overdose of the anesthetic, and the dorsal skinfold preparations were sacrificed for histological examination.

Histology and immunohistochemistry

For light microscopy (DM4000B, Leica Mikrosysteme, Wetzlar, Germany), 5 μm thick sections were cut and stained with hematoxylin and eosin (both Merck, Darmstadt, Germany). For immunohistochemical detection of CD31, for the presence of endothelial cells, formalin-fixed (Merck KgaA, Darmstadt, Germany) specimens of the dorsal skinfold chamber preparations were prepared according to standard procedures [23].

Statistics

The results are expressed as means ± SEMs. Differences between groups were assessed by one-way ANOVA, and differences within each group were analyzed by one-way repeated-measures ANOVA. To isolate overall differences, appropriate Student–Newman–Keuls or Dunn post hoc tests were performed. Differences were considered significant at p < 0.05.

Results

Microhemodynamics

Microhemodynamic parameters were constant and comparable in all experimental groups at any point in time. Values concerning venular diameters, volumetric blood flow, and wall shear rates were measured in postcapillary and collecting venules in the direct vicinity of the scaffolds immediately as well as 3, 6, 10, and 14 days after scaffold implantation. The diameters ranged from 20 to 40 µm. No significant differences could be observed in volumetric blood flow or wall shear rates between the groups throughout the entire experiment (Supplementary File 1).

Inflammatory response

The inflammatory response after scaffold implantation was assessed by monitoring the number of rolling and adherent leukocytes as well as microvascular leakage. The number of rolling leukocytes varied moderately in all experimental groups at any time point of the measurement. However, no significant change could be detected throughout the course of the experiment. On days 3, 6 and 10 after scaffold implantation, mice of all experimental groups showed a slight, but not significant, increase in the number of adherent leukocytes. Two weeks after implantation, the number of adherent leukocytes returned to the initial values as measured on day 0. The vascular permeability increased nominally in all experimental groups during the 14-day period. Overall, the implantation of scaffolds did not lead to significant changes in any of the measured inflammatory parameters throughout the 14-day observation period in either hypertensive or normotensive mice (Fig. 1).

Fig. 1
figure 1

Inflammatory response of the host tissue caused by the implantation of tissue engineering constructs. The number of rolling leukocytes, given as cells/min (a); the number of adherent leukocytes, given as cells/mm2 (b); and macromolecular leakage as an indicator of microvascular permeability (c) in postcapillary and collecting venules within the border zones of collagen-coated PLGA scaffolds (hypertensive mice: light gray bars; controls: dark gray bars), PLGA scaffolds additionally seeded with OLCs (hypertensive mice: light blue bars; controls: dark blue bars) or bmMSCs (hypertensive mice: light green bars; controls: dark green bars) immediately (day 0) as well as 3, 6, 10, and 14 days after implantation into the dorsal skinfold chamber (any group, n = 8). Data are presented as means ± SEMs

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Angiogenesis and neovascularization

In all experimental groups, typical signs of neovascularization could be detected for the first time on day 3 post implantation, characterized by the dilation of capillaries and the formation of capillary buds and sprouts, primarily originating from the venular segments of the striated muscle capillaries and postcapillary venules of the host tissue. At later time points, these sprouts protruded both from the border zone toward the center of the scaffold and from the center to the border zone, interconnecting with each other and building red-blood-cell-perfused vascular networks with increased functional capillary densities (Figs. 2 and 3). The increase in the microvascular networks was comparable and not significantly different between scaffold centers and border zones within the single groups at any point in time.

Fig. 2
figure 2

Microvascular networks 14 days after implantation. Intravital fluorescence microscopy of collagen-coated PLGA scaffolds (a hypertensive mice; d controls) and PLGA scaffolds additionally seeded with OLCs (b hypertensive mice; e controls) or bmMSCs (c hypertensive mice; f controls). Note the considerably more densely meshed microvascular networks in the control groups. Blue light epi-illumination with contrast enhancement by the addition of 5% FITC-labeled dextran 150,000. PLGA strands are marked with asterisks

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Fig. 3
figure 3

Time course of functional microvascular densities. Quantitative assessment of the functional microvascular density of newly formed microvessels, given in cm/cm2, of collagen-coated (Coll) PLGA scaffolds (hypertensive mice (BPH/2J): light gray bars; controls (BPN/3J): dark gray bars), PLGA scaffolds additionally seeded with OLCs (Coll + OLCs) (hypertensive mice: light blue bars; controls: dark blue bars) or bmMSCs (Coll + bmMSCs) (hypertensive mice: light green bars; controls: dark green bars) immediately (day 0) as well as 3, 6, 10, and 14 days after implantation into the dorsal skinfold chamber (each group, n = 8). Data are presented as means ± SEMs. *p < 0.05 versus controls with collagen-coated scaffolds and all hypertensive groups; °p < 0.05 versus hypertensive mice with collagen-coated scaffolds; +p < 0.05 versus the same group on the previous day; #p < 0.05 versus hypertensive mice with collagen-coated scaffolds

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Overall, quantitative analysis of functional capillary density demonstrated a continuous increase in microvascular density in all groups throughout the 14-day observation period. In that period, the most remarkable increase in newly formed capillaries occurred between days 3 and 6 after scaffold implantation. The growth rate of microvessels ranged between a 6-fold increase in the normotensive group equipped with bmMSC-seeded scaffolds to a 15-fold increase in the hypertensive group equipped with collagen-coated scaffolds.

Regarding hypertensive animals and healthy controls, separate quantitative analysis of functional capillary density demonstrated microvascular networks that were more densely meshed in the cell-seeded scaffolds than in the collagen-coated PLGA scaffolds. This trend persisted throughout the experiment, resulting in a significantly higher microvascular density within the cell-seeded scaffolds in the hypertensive or healthy animals than in the corresponding control groups on day 14. With sole regard to the cell-seeded scaffolds, in both groups (hypertensive and normotensive), there was no significant difference between the scaffolds that were seeded with OLCs and those that were seeded with bmMSCs (Figs. 2 and 3).

The growth pattern of the microvascular networks within the transplanted scaffolds was comparable in all mice and showed a steady increase from day 3 to day 14. However, in the hypertensive animals, the capillary density was generally reduced compared with the normotensive control groups, regardless of the type of transplanted scaffold (PLGAs coated only with collagen or seeded with OLCs or bmMSCs). This trend persisted throughout all examined time points, resulting in a significantly lower functional microvascular density in the hypertensive mice than in the normotensive controls at day 14 post scaffold implantation. The capillary densities within the cell-seeded scaffolds in hypertensive mice were approximately comparable to the microvascular networks induced by the scaffolds that were coated solely with collagen in healthy animals (Figs. 2 and 3).

Histology and immunohistochemistry

To confirm the intravital microscopic results, histology and immunohistochemistry were performed. Histological sections from paraffin-embedded specimens at day 14 showed clear differences in the abundance of microvascular structures depending on the animal group (hypertensive or normotensive) and the type of implanted scaffold (Fig. 4). In the collagen-coated scaffolds of hypertensive mice, microvascular structures were barely detected (Fig. 4g, h). The number of capillary-like structures in the cell-seeded scaffolds in hypertensive animals (Fig. 4i–l) was remarkably higher than that in scaffolds in the absence of seeded cells. The microvascular networks of the cell-seeded scaffolds in normotensive animals increased distinctly compared with those of all other scaffolds (Fig. 4c–f). Within these two groups, there was no significant difference between the scaffolds seeded with OLCs and those seeded with bmMSCs. The identity of the visible vascular structures as microvessels was confirmed by immunohistochemical detection of the endothelial cell marker CD31 (Fig. 5). The results of the immunohistochemical analysis were consistent with the results of the histological analysis.

Fig. 4
figure 4

Histological sections of paraffin-embedded specimens 14 days after implantation into the skinfold chamber. HE staining of collagen-coated PLGA scaffolds (controls: a and b; hypertensive mice: g and h) and PLGA scaffolds additionally seeded with OLCs (controls: c and d; hypertensive mice: i and j) or with bmMSCs (controls: e and f; hypertensive mice: k and l). Low-magnification pictures (left column: a, c, e, g, i, and k) provide an overview of the gross appearance of the scaffolds. After 14 days, the scaffold strands (asterisks) were embedded in the vascularized granulation tissue. The differences in microvascular densities are clarified by higher magnification pictures (right column: b, d, f, h, j, and l). Arrows denote the vascular structures

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Fig. 5
figure 5

Immunohistochemical detection of CD31 as a marker of endothelial cells (arrows). Collagen-coated PLGA scaffolds (a, b controls; g, h hypertensive mice), PLGA scaffolds additionally seeded with OLCs (c, d controls; i, j hypertensive mice) or with bmMSCs (e, f controls; k, l hypertensive mice). At day 14, a positive staining/signal for CD31 was detectable in all constructs, but marked differences in vascular density were observed depending on the composition of the constructs. PLGA strands are marked with asterisks. Left column (a, c, e, g, i, k): overview, right column (b, d, f, h, j, l): higher magnification

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Discussion

In the present study, we clearly demonstrated that cell seeding accelerates the vascularization of tissue engineering constructs in hypertensive mice. Nevertheless, neoangiogenesis in hypertensive BPH/2J mice was decreased compared to normotensive controls (BPN/3J mice) within the single groups throughout the entire 14-day observation period, resulting in significantly reduced capillary networks on day 14. This observation applied to the scaffolds seeded with organ-specific (OLCs) or undifferentiated progenitor cells (bmMSCs) as well as to the nonseeded scaffolds. The cell-seeded scaffolds showed a higher number of capillary-like structures within their particular group (hypertensive and normotensive) over the whole experiment than nonseeded scaffolds, with a significant difference on day 14. This result is in line with previous studies [23, 24, 29]. Microhemodynamic parameters were constant in any group and were comparable between single groups throughout the entire experiment.

Experimental hypertension can be induced in healthy mice by a variety of methods. Apart from surgically induced hypertension (e.g., by clamping the renal arteries), elevated blood pressure has also been generated by the injection of suitable bacteria, proteins, antibodies or phenol, leading to inflammation and kidney diseases [42,43,44,45]. The substitution of glucocorticoids and mineralocorticoids or appropriate influence of the renin-angiotensin system can also trigger hypertension [46]. Alternatively, a mouse strain characterized by genetic development of hypertension can be chosen. In the present study, we used the inbred mouse strain BPH/2J mainly for two reasons. First, we could prevent the animals from undergoing further invasive procedures. Second, the characteristics of this very common and well-studied hypertensive strain resemble those of primary hypertension in humans, since 90% of hypertensive patients suffer from this primary type [47, 48]. In primary hypertension, multiple genes contribute to the individual phenotype, each by diverse allele effects, penetrance, and contributions [46]. Therefore, no single genetic defect can explain the development of primary hypertension in humans. Using BPH/2J mice, we chose the phenotype-driven experimental approach, taking advantage of the natural variation among inbred strains to find quantitative characteristics in this disease model [46, 49]. Recent genetic analyses suggest a whole bandwidth of different genes that are responsible for the elevated blood pressure of BPH/2J mice [47, 48, 50]. Among others, genes regulating the balance of vasopressin, nitric oxide (NO), reactive oxygen species (ROS), the renin-angiotensin system, angiopoietin-like proteins, and histamine metabolism were identified [47, 48, 50].

For in vivo analysis of the early angiogenesis of tissue engineering constructs, the dorsal skinfold chamber is an excellent model [23,24,25, 28, 29, 51]. It allows repetitive intravital fluorescence microscopy for at least 14 days with subsequent histological and immunohistochemical analyses. We used PLGA scaffolds with a defined pore size of 250 µm, which have been demonstratedto be a well-suited three-dimensional basic material for tissue engineering constructs [23, 52]. These scaffolds in combination with intravital microscopy allowed us perfect comparability with previous investigations [2, 3, 23,24,25, 28, 29]. In contrast to other studies, we decided against the use of growth factors (e.g., vascular endothelial growth factor,VEGF) to support the vascularization of the transplanted scaffolds [2, 3]. Even in healthy organisms, the application of growth factors is extremely challenging. This applies not only to the dosage and combinations but also to a possible induction of uncontrolled vessel growth and neoplasms in unexpected regions of the organism [53, 54]. In combination with health risks such as hypertension, additional problems arise. Increased serum VEGF levels in association with increased carotid intima-media thickness were found in patients with metabolic syndrome [55]. Since hypertension is a main criterion of metabolic syndrome and elevated VEGF levels in hypertensive patients have been shown to correlate with cardiovascular risk, early microvascular and target organ damage, we decided against the usage of growth factors in the present study [56]. In this context, the risk of developing cardiovascular disease events exhibits a complex nonlinear relation to circulating VEGF concentrations, whereupon the underlying pathophysiological mechanisms remain to be elucidated [57].

To promote the vascularization of transplanted scaffolds, we seeded them with either bmMSCs or OLCs. For both cell types, in vitro tests of vascular differentiation capacities have been described in detail [9, 58,59,60]. In the current study, we decided to isolate and cultivate OLCs and bmMSCs from the healthy control strain (BPN/3J), which has the same genetic background as the hypertensive strain (BPH/2J) [61]. The reason for this is the commonly increased levels of ROS under hypertensive conditions. ROS create an inflammatory and oxidative microenvironment, which induces cell damage and the apoptosis of OLCs and bmMSCs [62]. Undifferentiated pluripotent bmMSCs are characterized by their ability to differentiate into smooth muscle cells, endothelial cells and hematopoietic supporting stroma [63, 64]. Due to these attributes, bmMSCs have the potential to support the development of a microvascular network. Very little, if any, is known about OLCs for tissue engineering applications, neither in hypertensive animal models nor in clinical studies. However, recently, the angiogenic effect of adipose-derived stem cells as a therapeutic target for enhancing the healing of hypertensive leg ulcers has been described [65]. In previous investigations, we studied the early angiogenesis of cell-seeded PLGA scaffolds in a mouse model of diabetes. Due to similar experimental procedures, these studies have excellent comparability, and we can conclude that seeding scaffolds with OLCs or bmMSCs could be a method to promote the vascularization of transplanted tissue engineering constructs even in disease models characterized by reduced angiogenetic potential [23, 29].

The scaffold implantation induced a slight inflammatory response in the host tissue, characterized by an increase in the number of rolling and adherent leukocytes and enhanced macromolecular leakage. This response was seen in all mice and is comparable to our earlier studies [3, 23, 29].

Since hypertension is associated with inflammation and an increase in ROS, a more distinct increase in the inflammatory parameters could have been expected in the BPH/2J mice [47]. NO biosynthetic and metabolic processes are decreased in BPH/2J mice, which may result in increased leukocyte adhesion to the endothelium [47, 66]. However, the preparation of the dorsal skinfold chamber and surgical scaffold implantation also led to inflammatory tissue damage and leukocyte activation [23, 52, 67]. Therefore, regarding the experimental conditions presented here, the surgically induced inflammatory host tissue response seems to have outweighed the increased leukocyte recruitment that may have been caused by reduced NO activity and elevated ROS levels in the hypertensive animals.

In tissue engineering, rapid blood vessel ingrowth into the transplanted constructs is the fundamental prerequisite for their successful incorporation. However, patients who would benefit from tissue engineering constructs often suffer extremely from widespread diseases that limit the capacity of neovascularization (e.g., hypertension, diabetes, obesity). According to our knowledge, in vivo studies about the angiogenesis of tissue engineering constructs in hypertensive organisms are extremely rare. As 25% of the global population suffers from elevated blood pressure, we paid special attention to the neoangiogenesis of the transplanted cell-seeded scaffolds in hypertensive organisms. Throughout the 14-day observation period, the development of microvascular networks in control groups was comparable to recent studies with the same experimental setup using (healthy) BALB/c and C57BL/6 mice [23, 29]. Therefore, we conclude that the increased angiogenesis in the cell-seeded control groups was caused by hypoxia-driven VEGF release from OLCs and bmMSCs. Regarding the hypertensive mice, microvascular networks within single groups showed a time progress that was very similar to that of healthy controls. Therefore, the increased vascularization in cell-seeded scaffolds in hypertensive mice was also most likely VEGF-driven based on the hypoxia of the OLCs and bmMSCs. However, reduced capillary densities were observed from day 3 onward in each case at any point of examination, resulting in significantly reduced capillary networks between the single hypertensive and normotensive groups on day 14. The possible mechanisms underlying the decreased neovascularization in hypertensive mice may be highly diverse. Recently, several genes have been detected that play key roles in the hypertension of BPH/2J mice [47, 48, 50]. Therefore, we would like to focus on these metabolic processes and their effects on neoangiogenesis in particular. Generally, impaired tissue perfusion under hypertensive conditions is caused by small arterial and arteriolar remodeling, microvascular rarefaction, and disturbed blood rheology [17]. In all of these alterations, ROS play an outstanding role as a cause as well as a consequence. Elevated blood pressure is associated with an increased circulation of ROS due to multiple mechanisms. In BPH/2J mice, elevated renal Ren1-gene messenger-RNA expression and lower levels of micro-RNA-181a, a negative regulator of renin messenger-RNA, lead to the activation of the renin-angiotensin system [48]. The consequence is an angiotensin-2-driven direct elevation of blood pressure and a boost in circulating ROS [17, 48]. Moreover, angiotensin-2 stimulates the expression of inflammatory mediators and adhesion molecules [17]. Different genes have been identified whose products have neurogenic blood-pressure-elevating effects, including the stimulation of inflammation and oxidative stress [47, 68, 69]. Oxidative stress is accompanied by an increase in ROS, leading to mitochondrial and nuclear DNA damage [47]. In this context, Marques et al. detected differential expression of various genes responsible for oxidative stress (e.g., Ddah1, Sod2, Me1, Nxn, Pink1, Arg2, Abcb8) in the hypothalamus of BPH/2J mice. Compared to BPN/3J mice, these different gene expressions result in a direct increase in ROS in BPH/2J mice [47]. Accordingly, elevated H2O2 levels could be detected in the aorta of BPH mice [31].

ROS and other inflammatory mediators affect blood rheology. Therefore, reduced fluidity is caused by negative changes in hematocrit, red blood cell deformability and aggregation, and leukocyte activation [22, 70]. In the present study, the average hematocrit elevation of BPH/2J mice was approximately 10% compared with that of normotensive BPN/3J mice. Such a discreet elevation does normally not lead to a change in blood rheology [71]. The inflammatory parameters (number of rolling and adherent leukocytes, vascular permeability) were comparable in all experimental groups (hypertensive and normotensive) during the 14-day observation period. Therefore, changes in volumetric blood flow and wall shear rate due to increased leukocyte activation were neither expectable nor detectable under the experimental conditions presented here. However, changes in the vessel structure and function of BPH/2 mice have been demonstrated. Baumbach and colleagues reported that BPH/2 mice exhibit cerebral arteriole hypertrophy but no reduction in external diameter [72, 73]. Additionally, BPH/2 mice were reported to have endothelial dysfunction in small-caliber arteries, demonstrated by a 25–50% reduction in maximal vasorelaxation in response to acetylcholine and bradykinin [72, 74]. Using Doppler ultrasound, BPH/2 mice have been shown to have impaired endothelium-dependent dilatation in femoral arteries [72, 75]. A comprehensive myographical examination of arteries from different vascular beds of BPH/2 and BPN/3 mice demonstrated that the arteries from the hypertensive mice have greater myogenic tone and impaired relaxation despite having similar passive qualities [72, 76].

Oxidative stress can also be induced by mitochondrial dysfunction and decreased NO production [47]. Since ROS lead to mitochondrial and nuclear DNA damage, a vicious cycle is the result [47]. In BPH/2J mice, elevated Dynll1-gene messenger-RNA was detected, which finally led to reduced NO bioavailability [47, 77, 78]. Decreased NO levels combined with ROS-induced NO inactivation are mainly responsible for the functional and structural microvascular rarefaction under hypertension [17]. Enhanced apoptotic endothelial cell death is also triggered by ROS under hypertensive conditions and contributes to rarefaction [17]. Problematically, hypertension and microvascular rarefaction can mutually be dependent. Rarefaction has been detected in individuals with a familial predisposition to hypertension, even if they themselves are normotensive [17, 79]. Mathematical modeling studies suggest that realistic degrees of rarefaction and remodeling can cause a significant increase in peripheral resistance and can amplify and stabilize an initial increase in blood flow or pressure, which leads to a vicious circle in hypertension [17, 80, 81].

Moreover, genetically induced upregulation of histamine, vasopressin, and angiopoietin-like proteins leads to a direct or indirect elevation of blood pressure and, therefore, to an intensification of the processes mentioned above [50, 82]. All these mechanisms result in reduced capillary recruitment and decreased blood flow reserve with decreased tissue perfusion [17]. Since decreased venular densities could also be detected under hypertensive conditions and because the very early vascularization of the transplanted tissue engineering constructs primarily originates from the venular segments of the striated muscle capillaries and postcapillary venules of the host tissue, the explanation of reduced capillary densities in the hypertensive groups is obvious [83].

The presented results highlight that the applied study design represents an excellent experimental setup for in vivo neovascularization studies of tissue engineering constructs under hypertensive conditions. Additionally, the existing comparability to similar examinations in healthy and impaired (diabetic) organisms is extremely valuable [2, 3, 23, 25, 26, 29]. Since most patients in whom tissue engineering constructs may be used are neither healthy nor young, studies under further metabolic changes (e.g., obesity or even advanced age) could follow. For hypertension, the challenge for the future will be to compensate for the decreased neovascularization in tissue engineering constructs. Therefore, two therapeutic targets can be considered: the tissue engineering construct and the patient. Concerning the graft, the prefabrication of vascular structures in vitro before transplantation seems to be a promising approach. For example, human microvascular endothelial cells (HMEC-1) or human umbilical vein endothelial cells (HUVECs) have been cultured under specific conditions to form capillary-like tube structures [84, 85]. Moreover, aortic fragments and capillary patterns have been seeded on specific scaffolds. Using appropriate culture systems, these vessel explants generate dense networks of tubular vascular sprouts [6, 7, 24]. Recently, a 2-week preincubation period of bmMSC-seeded PLGA scaffolds in Matrigel led to significantly accelerated angiogenesis after transplantation when compared with constructs that were not preincubated [25]. This positive effect on the neovascularization could even be enhanced by the coculture of aortic fragments and osteoblast-like cells for 14 days [28]. On the other hand, hypertensive patients should be a therapeutic target to enhance the chance of successful transplantation of tissue engineering constructs. Countless antihypertensives are in clinical use. Especially for angiotensin-converting enzyme inhibitors and AT(1) receptor blockers, the positive effects on angiogenesis have been shown. By activating bradykinin pathways, resulting in the generation of VEGF and NO, these drugs can reduce or even reverse microvascular rarefaction [86]. Furthermore, antioxidants offer enormous potential as they, e.g., reduce endothelial cell apoptosis and prevent the development of microvascular rarefaction when applied systemically during the growth of spontaneously hypertensive rats [87]. However, antihypertensive drugs and antioxidants could even enhance the quality of the tissue engineering construct. Recently, an accelerated decline in cardiac stem cell efficiency in hypertensive organisms has been shown [88]. Treatment with the antihypertensive metoprolol or with the antioxidant tempol increased cardiac stem cell migration and proliferation potential. Additionally, cellular senescence and oxidative stress in terms of intracellular ROS were reduced [89, 90]. Transferring the results of the present study into the clinic, autogenous cells would have to be isolated. Regarding this, the findings mentioned above could be very useful. The treatment of patients with metoprolol and/or tempol could restore stem cell efficiency before isolation. Moreover, the treatment of the isolated stem cells during cultivation could be considered. Finally, since an increasing number of genes are being detected that are responsible for impaired tissue perfusion under hypertensive conditions, individual gene therapy could be a hopeful option to enable tissue engineering in hypertensive patients [48].