Oxygen-releasing biomaterials
Steg, Hilde
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Publication date: 2018
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5
Biocompatibility and proof of
concept of novel
oxygen-delivering microspheres
Arina T Buizer, Hilde Steg, Sjoerd K Bulstra, Albert G Veldhuizen, Dirk W Grijpma, Willy de Haan-Visser, Willem Woudstra, Roel Kuijer
Manuscript in preparation
Abstract
Tissue repair often needs to take place in poorly vascularised areas. Due to insufficient removal of waste products and decreased supply of nutrients and oxygen many repair cells die before they contribute to new tissue formation. To support cell survival until a sustained oxygen supply has been accomplished, oxygen-delivering microspheres based on poly(trimethylene carbonate) (PTMC) and calcium peroxide were prepared and evaluated. Cytotoxicity of these microspheres was assessed in vitro by culturing L929 cells with a microsphere extract and subsequently analyzing cell metabolic activity. The microspheres were tested in vivo by implanting them in subcutaneous pockets in mice for 1 and 6 weeks, followed by histological assessment of local tissue reactions. As a proof of concept, oxygen-releasing microspheres were implanted in a random pattern in a devascularized skin flap in 12 mice. Photographs were taken of the skin flaps at 3, 7, and 10 days after surgery, and skin necrosis was assessed by three independent observers using Image J software. Histologic examination of the skin flaps was done after termination of the animals at day ten. In in vivo tests, the oxygen-delivering microspheres were shown to be biocompatible. Significantly less skin necrosis was observed in skin flaps under which oxygen-releasing microspheres were implanted than in skin flaps under which non-oxygen-releasing microspheres were implanted at day 3, 7, and 10 after surgery. This finding suggests that these oxygen-releasing composite microspheres support tissue survival under ischemic circumstances.
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Introduction
For adequate tissue repair under ischemic circumstances the accomplishment of an adequate oxygen supply to repair cells remains a big challenge1. Within the body, oxygen is transported efficiently via the extensive
vascular network that the body possesses2. Given that the maximum diffusion
distance of oxygen within the body is around 100-200µm, the vascular network must be extensively branched to bridge this maximum oxygen-diffusion distance3,4. In ischemic tissues vascularization is often inadequate. Oxygen
provision to repair cells is thus compromised1,3,5. The ischemic conditions are an
important cause of death of repair cells and they are an important contributor to the limited success rate of ischemic tissue recovery.
Several methods aiming at improvement of local vascularization have been proposed to increase repair-cell survival under ischemic circumstances. A frequently studied option is the application of precursor cells, which are cells that have the capacity to differentiate into the desired type of tissue. An example is the application of adipose-derived mesenchymal stem cells (MSCs) in heart muscle tissue after myocardial infarction6. To improve the angiogenic potential of
progenitor cells, cells may be exposed to hypoxic circumstances prior to implantation, in order to increase the production of factors that increase vascular ingrowth7,8. This process is called hypoxic preconditioning. A different option to
increase vascular ingrowth is the application of exogenic angiogenic growth factors (AGF), which stimulate the ingrowth of blood vessels9. Also, the
application of cells that are modified prior to implantation so that they produce more AGF has been investigated10. In some situations, application of cells alone
may not be sufficient for tissue repair. A scaffold may be needed as a matrix for cells to grow upon. To improve vascularization in cell-scaffold complexes many techniques have been studied. Examples of these techniques include co-seeding of endothelial cells combined with several types of precursor cells on a scaffold11–13
and the application of (AGF) in or near a scaffold14–16.The results were variable. A
promising option to increase repair cell survival is to supply repair cells with oxygen from an external source, so that they can survive until vascular ingrowth has occurred and thus sustained oxygen supply has been accomplished.
Oxygen-release systems consisting of a polymer carrier material combined with a peroxide have been created by several research groups17–19. These usually
comprise a hydrolysable polymer combined with a peroxide material. In this study, a composite consisting of poly(trimethylene carbonate) (PTMC) as the polymer carrier component and calcium peroxide (CaO2) as oxygen-donating
component, was evaluated. PTMC was chosen as it is degraded by means of surface erosion without production of acidic waste products. CaO2 was chosen
for its favorable oxygen-release profile combined with its availability in high purity20. Whenever CaO2 comes into contact with water, the following chemical
reactions occur, resulting in the release of oxygen (equations 1 and 2): 𝐶𝑎𝑂2+ 2𝐻2𝑂 ⇌ 𝐶𝑎(𝑂𝐻)2+ 𝐻2𝑂2 [Equation 1]
2𝐻2𝑂2 ⇌ 2𝐻2𝑂 + 𝑂2(Catalase) [Equation 2]
A slow release system of oxygen is created by incorporating the peroxide into the PTMC polymer matrix. As the polymer is degraded by surface erosion, the peroxide is gradually exposed to water and oxygen is released gradually. The PTMC-calcium peroxide (PTMC/CaO2) composite will thus function as a
slow-release system for oxygen. By adjusting the dosage of the microspheres, the type of polymer, or the concentration of peroxide, the oxygen-release can be adapted to the oxygen demand. For application in bone, an important factor is the mechanical strength of the material. PTMC is an elastic polymer with limited mechanical properties for application in loaded bone. Therefore, microspheres were produced that can be combined with porous bone replacement materials so that sufficient mechanical support is provided. The microspheres produce oxygen for several weeks in vitro21.
To take a first step towards application of this material in clinical practice, biocompatibility of the PTMC/CaO2 composite was tested in vitro using an extract
test and in vitro using a subcutaneous pocket model in mice. As a proof of concept of the working potential of these microspheres, oxygen-releasing PTMC/CaO2 microspheres were implanted under a random pattern
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Materials and methods
Biomaterial manufacture
The PTMC/CaO2 composite microspheres were manufactured according
to the process described by Steg et al21. Briefly, microspheres were prepared
from a suspension of 5% (w/w) CaO2 (75% purity, sieved to particle sizes <74µm,
Sigma-Aldrich, Steinheim, Germany) in a solution of 3.5 % (w/v) of poly (1,3-Trimethylene Carbonate) (PTMC, Mn=220 kg/mo) in acetonitrile (Merck, Darmstadt, Germany) using an oil-in-oil solvent evaporation method. As a control, PTMC microspheres not containing CaO2 were produced using the same
manufacturing process. The spheres were stored at -20°C until use.
In vitro biocompatibility testing
In preparation of the experiment, extracts of PTMC and PTMC-CaO2
microspheres were made by adding 15mg of either type of microspheres to one milliliter of standard medium, consisting of DMEM high glucose (Invitrogen, Paisley, UK) supplemented with 10% fetal bovine serum (FBS, Invitrogen, Paisley, UK) and 1% Glutamax (Life technologies, Eugene, OR, USA). Subsequently, the suspension was incubated for 24 hours in a shaking water bath at 37°C. After 24 hours, the extract was centrifuged at top speed for five minutes. Concentrated extracts (100%) and dilutions of 75%, 50% and 25% were made. A positive control was made by dissolving 1% sodium lauryl sulfate (SLS, Sigma-Aldrich, Steinheim, Germany) in medium. All extracts and the positive control were prepared in triplicate.
L929 mouse fibroblast cells (Biosorb, the Netherlands) at passage 47 were seeded at a density of 10,000 cells per well in a 96 well plate (Greiner Bio-one, Alphen aan den Rijn, the Netherlands) and were allowed to adhere at 37°C and at 5% CO2 for 24 hours. 12 wells of each plate were filled with standard medium
only, to serve as a blank. Then, the medium was replaced with 100µl of extract. Each dilution of each extract was added to five wells. Subsequently, the cells were incubated at 37°C and at 5% CO2 for 24 hours. The MTT assay was performed
bromide / Thiazolyl Blue Tetrazolium Bromide (MTT) (Sigma-Aldrich, Steinheim, Germany) in full culture medium (MTT medium), replacing the medium of the cells with 50µl MTT medium, and incubating the plate at 37°C for two hours in the dark. After two hours, MTT medium was carefully removed from the wells and 100µl of 2-propanol (Merck, Darmstadt, Germany) was added to each well. The plates were shaken for two minutes to dissolve the produced formazan, after which absorbance was read at 570nm using a plate reader (Fluostar Optima, BMG labtech, Olfenburg, Germany). Background absorbance was read at 650nm. Prior to viability calculations, background absorbance values were subtracted from the absorbance values read at 570nm. The viability of the cells was calculated using the following formula:
𝑉𝑖𝑎𝑏𝑖𝑙𝑖𝑡𝑦 % = 𝐴𝑏𝑠𝑜𝑟𝑏𝑎𝑛𝑐𝑒 𝑜𝑓 𝑛𝑒𝑔𝑎𝑡𝑖𝑣𝑒 𝑐𝑜𝑛𝑡𝑟𝑜𝑙 𝐴𝑏𝑠𝑜𝑟𝑏𝑎𝑛𝑐𝑒 𝑜𝑓 𝑒𝑥𝑡𝑟𝑎𝑐𝑡 𝑥 100% [Equation 3] All experiments were performed in triplicate.
Animals and surgical procedure
The Experimental Animal Committee of the University Medical Center Groningen approved all animal experiments, and Dutch national guidelines for animal care were followed.
In vivo biocompatibility testing
Twelve female Balb/c Ola/Hsd (Harlan, Horst, the Netherlands) mice of 7 weeks old were used for biocompatibility tests. The operations were carried out under aseptic conditions. General anesthesia was induced using isoflurane 4% and maintained using isoflurane 2%. Animals were administered buprenorphine 0.1mg/kg sc once immediately before surgery. Two six millimeters long incisions were made on the backs of the mice on the midline and running parallel to the spine. Both incisions were one centimeter apart, the most cranial starting right caudal of the scapulae. From each incision, two subcutaneous pockets were created using blunt dissection, one left lateral to the incision and one right lateral to the incision. The pockets were ten millimeters deep. On the bottom of each pocket, 25mg of microspheres was placed. In each pair of pockets originating at
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one incision, one dose of PTMC microspheres was implanted on one side andone dose of PTMC/CaO2 microspheres was implanted on the opposite side. The
type of material that was implanted on the left side of the incision was randomized. The wounds were closed using resorbable sutures. Half of the animals were terminated at one week after implantation of the biomaterials and half of the mice were terminated at six weeks after implantation by cervical dislocation. The biomaterial and near surroundings were excised after termination.
Skin flap test
For this experiment, the model used by Harrison et al17 was adapted to
our needs. Twelve female BALB/c mice (BALB/c OlaHsd, Harlan, Horst, the Netherlands) of 6-8 weeks old were randomly divided in a control group of 6 mice receiving PTMC microspheres and an intervention group of 6 mice receiving PTMC/CaO2 microspheres. The operative procedure was performed under
anesthesia using isoflurane 2%. The animals were shaved and subsequently the stubbles were removed using depilation cream. A cranially based skin flap was created by making two incisions of three centimeters long running parallel to the spine, 0.5cm of the midline of the animal. Both incisions were connected with a transverse one-centimeter long incision located at the caudal end of the longitudinal incisions. The skin was bluntly dissected from the muscular layer. Care was taken that no large vessels were included in the skin flap, so that blood supply would be limited to the cranial base of the flap. Then one longitudinal incision and the transverse incision were sutured using Monocryl 5.0 (Ethicon, Norderstedt, Germany) and interrupted sutures. 100 milligrams of microspheres were applied on the muscular layer on the most caudal 2x1 cm area under the skin flap and spread evenly. The second longitudinal incision was sutured as well. Carprofen 5mg/kg sc once per 24 hours was administered routinely under anesthesia using isoflurane 2% for the first three days after surgery. The animals had access to food and water ad libitum and were housed in pairs in standard cages. Ten days after surgery the animals were terminated by cervical dislocation under general anesthesia. The skin flaps were excised in a standard manner and further processed for histological examination.
Processing of tissue samples
The tissue samples acquired in the biocompatibility tests were fixated in paraformaldehyde 3.7% (Boom, Meppel, the Netherlands) and cut in half after fixation. One half was embedded in paraffin and the other half was embedded in 2-hydroxyethyl methacrylate (HEMA) (Technovit® 8100, Heraeus-Kulzer, Wehrheim, Germany) according to the manufacturer’s instructions. A microtome was used to cut paraffin sections of 5µm and HEMA sections of 4µm. Three paraffin embedded slides were stained with Massons trichrome stain (Accustain® trichrome stains (Masson), Sigma-Aldrich, Egham, UK) according to the manufacturer’s instructions. Three paraffin embedded slides were used for immunohistochemical staining with a CD3-antibody for identification of lymphocytes and with antibody F4/80 for identification of macrophages. The slides were dewaxed in xylene and rehydrated using phosphate buffered saline (PBS). The slides were preincubated with PBS supplemented with 10% serum from the species that produced the secondary antibody. Subsequently they were incubated with either rabbit-anti mouse CD3 monoclonal primary antibody (Abcam, Cambridge, United Kingdom) in a 1:500 dilution or rat-anti mouse F4/80 monoclonal primary antibody (BioRad, Veenendaal, the Netherlands) in a 1:1000 dilution. The slides were washed with PBS and endogenous peroxidases were blocked using H2O2. After washing the slides with PBS, the secondary antibodies,
goat-anti rabbit-HRP (Agilent, Middelburg, the Netherlands) (dilution 1: 100) and goat-anti rat-HRP (BioRad, Veenendaal, the Netherlands) (dilution 1:200) respectively, were applied. Subsequently, the slides were washed with PBS and incubated with 3,3’-diaminobenzidine. After washing with PBS, the slides were counter-stained with hematoxylin (Merck, Darmstadt, Germany).
Three plastic embedded slides were stained with hematoxylin and eosin (Merck, Darmstadt, Germany). Quantification of the foreign body reaction was done according to an adapted system as proposed by Cohen22. All slides were
photographed at a 400x magnification and the numbers of polymorphonuclear cells, plasma cells and giant cells and the amount of tissue necrosis and fibrosis were assessed by 2 independent observers. The results were recorded in tables as indicated in tables 1 and 2 (modified from ISO 10993-6: 2007). Three parts of each tissue sample were thus assessed by 2 observers. The numbers of
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lymphocytes and macrophages were counted using an automatic cell countingcomputer program. A score of the intervention tissue after subtraction of the control sample scores of 0.0 up to 2.9 was considered non-irritant, a score of 3.0 up to 8.9 was considered slightly irritant, a score of 9.0 up to 14.9 was considered moderately irritant and a score of ≥15 was considered severely irritant.
Photography and image analysis in skin flap experiments
At days 3, 7 and 10 after the surgery, the animals were anaesthetized using isoflurane 2% via a non-rebreathing face mask. The skin flap on their back was photographed using a digital camera and standard lighting. A ruler was included in each picture for calibration purposes. The area of brown discoloration due to skin necrosis was assessed using Image J analysis software by three independent observers blinded for the applied treatment. Each skin flap was assessed trice by each observer. The extent of skin necrosis was expressed in percentage of the skin flap area that showed necrosis.
Processing of skin flaps in skin flap experiments
The skin flaps were cut in 4 equally sized longitudinal strips after excision from the animals and fixated in paraformaldehyde 3.7% (Boom, Meppel, the Netherlands). The strips were washed, dehydrated and then embedded in Technovit® 8100 (Heraeus-Kulzer, Wehrheim, Germany). Four µm thick sections were cut using a microtome. The sections were mounted on Superfrost slides (Thermo Scientific, Braunschweig, Germany) and stained with hematoxylin (Merck, Darmstadt, Germany) and eosin (Merck, Darmstadt, Germany). Images were studied using a DMR microscope (Leica HC, Wetzlar, Germany) equipped with a Leica DFC 420C camera (Leica, Wetzlar, Germany).
Statistical evaluation
The Statistical Package for Social Sciences (SPSS) was used for statistical evaluation of the results. Distribution of the data was assessed using a Shapiro-Wilk test. P values were calculated using a Mann-Whitney U test for not-normally
distributed data (skin necrosis data) and a T-test was used for normally distributed data (all biocompatibility data). A p value of >0.05 was considered to be significant.
Results
In vitro biocompatibility testing
According to ISO standard 10993-5, an agent that causes the viability percentage in cytotoxicity assays to be lower than 70%, is considered to have cytotoxic potential. Cell viability percentages after in vitro cytotoxicity tests are indicated in Table 4. At all tested conditions, cell viability was significantly lower than when cells were cultured with standard medium. When comparing cell viability of cells cultured with PTMC or with PTMC/CaO2 microspheres extract,
cell viability was significantly lower in cells cultured with PTMC/CaO2
microspheres extract in 75% and 50% dilution, but not at other dilution. All extract dilutions of PTMC microsphere extract resulted in cell viabilities higher than 70%, so PTMC microspheres are not cytotoxic at all tested concentrations of the extract Figure 1. After addition of 100% and 75% PTMC/CaO2
microspheres extract to the cells, cell viability was reduced to 62% and 65%, respectively. These composite microspheres thus have some cytotoxic effect in vitro. At lower dilutions of PTMC/CaO2 microspheres extract, a cytotoxic effect
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Figure 1: Bar chart indicating the absorbance measured after MTT test in the cytotoxicity assay. As a negative control, cells that were grown without extract were used. SLS 10 gr/liter was used as a positive control. Error bars indicate standard deviation.
Figure 2: Bar chart indicating the average biocompatibility scores, given as described in the materials and methods section. There was no significant difference between the PTMC and the PTMC/CaO2 groups at both time points. Error bars indicate standard deviation.
0 0,2 0,4 0,6 0,8 1 1,2 Abs orbanc e Test condition PTMC PTMC-CaO2 0 5 10 15 20 25 30 35 Week 1 Week 6 Av erage bi oc ompati bi lity s c ore
Number of weeks after implantation
PTMC PTMC-CaO2
In vivo biocompatibility testing
AnimalsNo complications occurred during surgery or in the postoperative period. No preliminary termination of animals was needed. Animal discomfort was estimated to be 3/6, with 1 indicating slight discomfort and 6 indicating serious discomfort.
Biocompatibility testing
Tissue samples of subcutaneous pockets containing either PTMC or PTMC/CaO2 microspheres were assessed for biocompatibility at 1 and at 6
weeks after implantation according to Table 1 and Table 2. The difference in biocompatibility scores between PTMC and PTMC/CaO2 microspheres was -0.8
at one week and 0.6 at six weeks (Figure 2). This indicates that the PTMC/CaO2
microspheres are non-irritants. The difference in biocompatibility scores was not significant at both time points.
In Figure 3, representative pictures of histological specimens of the implants using four different stainings are shown. In all pictures, thin capsules surrounding the implants can be identified. These capsules are fibrotic as shown through the Masson’s trichrome stain. The F4/80 immunostaining shows these capsules contain many macrophages. On the hematoxylin-eosin stained slides only few foreign body giant cells could be identified, both in PTMC implants as in PTMC/CaO2 implants. The numbers of polymorphonuclear cells, plasma cells and
giant cells and the amount of necrosis are low in both types of material tested. The amount of fibrosis is somewhat higher, but still there is no difference between both groups. The numbers of macrophages and lymphocytes are larger, but again, there is no significant difference between both tested materials (Table 3).
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Figure 3: Representative pictures of histological specimens of PTMC (left column) and PTMC/CaO2 microspheres (right column) stained with hematoxylin-eosin (a and b), Masson’s trichrome (c and d), immunostaining for macrophages using F4/80 macrophage antibody (e and f) and for lymphocytes using CD3 antibody (g and h) at magnification 400x. Scale bars indicate 50μm.
Skin flap tests
AnimalsAll animals tolerated the operations well and no complications occurred. There were no early dropouts. Animal discomfort was estimated to be 3/6. Skin necrosis
Comparison of the three independent evaluators of the amount of skin necrosis, revealed that the inter-rater reliability had an ICC of 0.803 (95% CI: 0.599-0.890). The intra-rater reliability had ICCs of 0.983, 0.971, and 0.996 for raters 1, 2, and 3 respectively. The amount of necrosis was variable within both the PTMC group and the PTMC/CaO2 group. Therefore, the results were not
normally distributed; medians are given in Table 5. At 3-, 7- and 10-days post-surgery skin necrosis was significantly higher in the PTMC group than in the PTMC/CaO2 group. These results indicate that the PTMC/CaO2 microspheres
did support cells under circumstances of disturbed vascularization in contrast to PTMC microspheres not releasing oxygen.
Cell type/response Score
0 1 2 3 4
Polymorphonuclear cells 0 Rare, 1-5 phf* 5-10/phf Heavy infiltrate Packed Plasma cells 0 Rare, 1-5 phf 5-10/phf Heavy infiltrate Packed Giant cells 0 Rare, 1-5 phf 5-10/phf Heavy infiltrate Packed Necrosis 0 Minimal Mild Moderate Severe Lymphocytes 0 Rare, 1-5 phf 5-10/phf Heavy infiltrate Packed Macrophages 0 Rare, 1-5 phf 5-10/phf Heavy infiltrate Packed Fibrosis 0 Narrow band Moderately Thick band Extensive
thick band band
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Biomaterial ID:
Test sample Control sample
Slide type Scoring item A B C A B C
HE Polymorphonuclear Cells Plasma cells Giant cells Necrosis CD3 stain Lymphocytes F4/80 stain Macrophages Sub-total (x2)
Massons trichrome Fibrosis
Subtotal Total
Test(-) control =
Table 2: Score form histologic evaluation of implants
Cell type/response Average score per type of microspheres
PTMC PTMC/CaO2 PTMC PTMC/CaO2 wk 1 wk 1 wk 6 wk 6 Polymorphonuclear cells 1.4 (0.6) 1.4 (0.6) 1.1 (0.6) 1.1 (0.6) Plasma cells 1.3 (0.6) 1.4 (0.6) 0.8 (0.5) 0.9 (0.7) Giant cells 0.6 (0.6) 0.7 (0.7) 0.8 (0.7) 0.7 (0.7) Necrosis 0.9 (0.8) 0.7 (0.6) 0.9 (0.6) 0.8 (0.6) Lymphocytes 2.9 (0.4) 2.2 (0.9)* 2.8 (0.6) 2.8 (0.4) Macrophages 3.8 (0.4) 3.7 (0.5) 3.8 (0.4) 3.9 (0.4) Fibrosis 1.8 (1.0) 2.0 (1.1)* 1.6 (0.9) 1.9 (0.9)
Table 3: Average histology scores per cell type and per type of material. Standard deviations are indicated in brackets. A significant difference in scores between PTMC and PTMC/CaO2 microspheres is indicated with an asterisk.
Extract 100% Extract 75% Extract 50% Extract 25% Positive control
PTMC 78 73 78 83 0
PTMC/CaO2 62 65 80 91 0
Table 4:Viability percentages of cells after in vitro cytotoxicity tests.
Day 3 Day 7 Day 10
PTMC PTMC/CaO2 PTMC PTMC/CaO2 PTMC PTMC/CaO2
Median 74.0 45.0 73.5 62.5 86.0 77.5
Table 5: Median percentage of skin necrosis at three time points in mice with PTMC microspheres implanted and in mice with PTMC/CaO2 microspheres implanted.
Figure 4: Representative pictures of random pattern devascularised skin flap after implantation of microspheres. Under the skin flap in the mouse on pictures a, c, and e, PTMC microspheres were implanted. Under the skin flap in the mouse on pictures b, d, and f, PTMC/CaO2 microspheres were implanted. Pictures a and b were taken 3 days after surgery, pictures c and d were taken 7 days after surgery and pictures e and f were taken 10 days after surgery.
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Histology
Gross histologic examination at a low magnification (25x) gave an indication of the course of skin necrosis along the full length of the skin flaps. At the cranial end of the skin flaps usually morphologically normal skin tissue was visible (see Figure 5), with the characteristic dark colored, several cells thick, epidermal layer and the presence of typical papillary structures. Moving along to the caudal end of the skin flap, the tissue morphology changed. The epidermal layer became thinner or even disappeared, and tissue architecture became less organized. The papillary structure of the epidermal layer became less evident or disappeared as well. At the caudal end of the skin flaps often a recurrence of normal skin architecture could be observed, as the skin flaps were excised including a rim of healthy skin tissue around the skin flaps. The histologic specimens shown in Figure 5 are representative of the difference in tissue necrosis between control and experimental group skin flaps.
Figure 5: Histologic specimens of skin flaps, HE staining. The specimens in figure a were taken from a skin flap under which PTMC microspheres were implanted. The specimens in figure b were taken from a skin flap under which PTMC/CaO2 microspheres were implanted. The irregular shaped figures are pictures of the full skin flap at a magnification of 25x. The rectangular pictures represent detailed pictures of the skin tissue taken at a magnification of 400x. Scale bars in the rectangular pictures represent 50µm. In the pictures of normal skin, the stratum corneum is indicated with a black arrow, the stratum granulosum is indicated with a white arrow and the stratum spinosum is indicated with a black asterisk.
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Histologic examination of the skin flaps at a higher magnification (400x)showed intact skin tissue at the cranial site of the skin flaps, with an epidermal layer including a clear stratum corneum, stratum granulosum and stratum spinosum. The cells were intact, and the cell nucleus was clearly visible. When proceeding to the more caudal part of the skin flap, the tissue architecture became disorganized, and the laminar structure of the skin could not always be recognized. The eosinophilia of the cytoplasm of the cells was striking and the cell nuclei were less sharply defined or could not be identified anymore. The combination of eosinophilia and deterioration of the cell nuclei is a clear indication that the cells were necrotic. At the cranial end of the skin flap, the tissue architecture appeared healthy.
Discussion and conclusion
The aim of this study was to assess the biocompatibility of oxygen-releasing composite PTMC/CaO2 microspheres and to explore the effect of implanted
oxygen-releasing microspheres on the process of necrosis in partly devascularized skin flaps. In in vitro tests, the composite microspheres were biocompatible at 50% or lower dilutions of the composite microsphere extract. In in vivo tests, the oxygen-delivering microspheres were shown to be biocompatible. In comparison to PTMC microspheres, subcutaneous implantation of oxygen-releasing PTMC microspheres resulted in a significant reduction of the amount of necrotic skin tissue at all three follow-up moments. These findings suggest that the release of oxygen from the PTMC/CaO2 microspheres may
support survival of the otherwise ischemic skin cells and thus may aid in the prevention of necrosis. In the most cranial parts of the skin flaps, the skin morphology was still intact 10 days after implantation of the microspheres. In the caudal regions, skin necrosis was clearly visible. These results suggest that there is an ischemic gradient within the skin flap, the most caudal area being the most ischemic, while the most cranial area is less or not ischemic.
The lower cell viability of L929 cells upon exposure to 100% and 75% concentrations of the PTMC/CaO2 microsphere extract may be explained by the
of calcium peroxide to water, hydrogen peroxide is formed, as was shown in equation 120. A burst release of oxygen from PTMC/CaO2 microspheres upon
exposure to water was shown by Steg et al21. Hydrogen peroxide is an
intermediate product in the eventual reaction of calcium peroxide with water to produce oxygen. It could be that cytotoxic concentrations of hydrogen peroxide are present in the 100% and 75% concentrations of the PTMC/CaO2 microsphere
extract, thus causing higher cell death than at lower concentrations of the extract. No hydrogen peroxide is produced during the preparation of the PTMC microsphere extract, which supports the hypothesis that the cytotoxic effect of high concentrations of PTMC/CaO2 microsphere extract is caused by excess
hydrogen peroxide.
Based on these test results, no indication of the amount of microspheres that needs to be used in clinical practice can be given. A dose response curve should be created, preferably in in vivo tests, to be able to calculate the amount of microspheres that should be applied for adequate enhancement of tissue regeneration. For this experiment, extracts of 15 micrograms of microspheres in one milliliter of medium were used. However, it is expected that in clinical practice dosages of 15 mg of microspheres per milliliter volume of tissue is too high to be used. The volume of the microspheres would in that case not allow adequate dispersion throughout the tissue. It is expected therefore, that the cytotoxic effect of the PTMC/CaO2 microsphere extract at high concentrations is
not clinically relevant.
PTMC is a biocompatible polymer, which makes it a suitable blank to test the biocompatibility of PTMC/CaO2 microspheres23–25. The polymer is degraded
enzymatically in the presence of lipase or cholesterol esterase26,27. Macrophages
produce cholesterol esterase24,28. PTMC degradation occurs through a surface
erosion process23–26. In in vitro tests, it was shown that when macrophages were
cultured in the presence of PTMC films but separated from the films by a semi-permeable membrane, no degradation of the PTMC films occurred. However, when the macrophages were cultured after being adhered to PTMC films, the material was eroded. Culturing fibroblasts on PTMC films did not result in PTMC degradation28. These results suggest that macrophage contact with PTMC is
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in vivo, initially, a fibrin layer is formed around the implant. In this layer,macrophages and giant cells can be found. A couple of days after implantation, a fibrous capsule containing macrophages and foreign body giant cells can be identified23,24,29. The tissue reaction 5-7 days after implantation remains
discordant, maybe due to different implantation sites or differences between animal species that were used in different studies. Some authors describe layers of foreign body giant cells and macrophages containing phagocytosed PTMC particles around the implant. In the 2 to 52 weeks after implantation the material weight and volume decreases23,24,29. Others describe only multilayered fibrous
capsules around the implants, with only few macrophages or nodules of foreign body giant cells25,27. As well around the PTMC microspheres as around the
PTMC/CaO2 microspheres, we found large macrophage infiltrates, which
corroborates with the findings of Pego and Bat23,24,29. This supports the
assumption that PTMC is degraded by macrophages when in contact with the polymer28.
To our knowledge, oxygen-releasing biomaterials prepared from a polymer and a peroxide salt as oxygen donor have been tested in vivo only once. Harrison et al.17 implanted films made out of a composite of Poly(D,L-lactide-co-glycolide)
(PLGA) and sodium percarbonate under random pattern devascularized flaps in mice and found that two and three days after implantation of the film, skin necrosis was significantly less in flaps under which oxygen-releasing PLGA films were implanted than in skin flaps under which control films not releasing oxygen, were implanted. Seven days after implantation of the PLGA films, the amount of skin necrosis was similar in oxygen-releasing films and films that did not release oxygen. The PTMC/CaO2 microspheres that were tested in our study slowed
down the occurrence of necrosis in a devascularized skin flap for a longer period, and even after ten days, skin necrosis was significantly lower after implantation of oxygen-releasing microspheres. An explanation for the shorter working time of the PLGA-sodium percarbonate composite could be the difference in degradation mechanism between PTMC and PLGA. PTMC is a polymer that degrades by surface erosion28,30, in contrast to PLGA, which degrades by bulk degradation31.
Surface erosion entails the breakdown of a material in a layer-by-layer mode, so that the surface of the polymer degrades first, and the core of the material last. In
bulk degradation, water penetrates the polymer matrix and the polymer chains break down by hydrolysis31. It is likely, that the surface erosion process, as is
applicable to PTMC, enables a more gradual and prolonged release of oxygen from the polymer carrier than does bulk degradation, as is applicable to PLGA. The influence of the type of carrier polymer on oxygen-release from a polymer-peroxide composite is confirmed by Pedraza et al.19, who produced a
polydimethylsiloxane (PDMS)-calcium peroxide composite that released oxygen for up to four weeks in vitro. Pancreatic islet cells grown under hypoxic circumstances in the presence of PDMS-calcium peroxide disks had higher metabolic activity, higher insulin production, less lactate dehydrogenase release and lower caspase activity than pancreatic islet cells grown without an oxygen-releasing disk. However, PDMS shows hardly any degradation in vivo, a material property that is not always desirable in in vivo application19.
Although oxygen-releasing microspheres were implanted under the skin flap in this study, still skin necrosis occurred after a couple of days. Exhaustion of the oxygen supply may be a cause of the skin necrosis. It is also possible that not a lack of oxygen was the cause of the skin necrosis, but that a shortage of other essential nutrients that would otherwise be supplied via blood-borne transport, such as proteins or vitamins.
The possible applications of oxygen-delivering biomaterials are manifold. Plastic surgeons frequently use skin flaps in clinical practice. A frequently occurring complication is necrosis of a skin flap32. As has been shown in this
study, oxygen-releasing microspheres may aid in the prevention of necrosis in a skin flap. The use of oxygen-releasing materials has also been proposed for supporting regeneration of cardiac tissue, for example after myocardial infarction33,34. After myocardial infarction, part of the heart muscle tissue is
damaged and needs to be regenerated to regain optimal heart function. Regeneration of cardiac tissue by stem cell therapy has been inefficient until now. One of the leading causes of this inefficacy is cell death of the cells applied in heart tissue due to ischemia34. Oxygen-delivering biomaterials may aid cell
survival and may have the potential to make heart tissue regeneration more successful. Other possible applications of oxygen producing biomaterials can be in
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regeneration of bone tissue in maxillofacial or orthopedic surgery, for treatmentof large ischemic ulcers, and several other applications.
Assuming that vascular ingrowth takes place at an ingrowth rate of 0.5mm per day35, it would take about 10 days to revascularize one centimeter of tissue,
whenever blood vessels can grow in from two opposite sides. Repair cells should thus survive for time periods of up to several weeks until vascular ingrowth is completed, which means that oxygen-delivering scaffold materials should provide oxygen for weeks as well. In vitro, PTMC/CaO2 microspheres produced oxygen
for about 3 weeks21. In vivo however, the oxygen-releasing composite
microspheres had a significant positive effect until at least ten days after implantation of the material. It is yet unknown how long the oxygen-release keeps on supporting the skin cells and thus aids in preventing necrosis in the animal model that was used, but ten days are a good start. Perhaps the oxygen-release from this type of materials could even be lengthened, by using a densely cross-linked PTMC material, which is degraded slower and thus a prolonged oxygen-release may be accomplished. Using an even more hydrophobic carrier polymer, which is degraded more slowly in the body, may also lengthen oxygen-release from a polymer-peroxide construct. Adjustment of the peroxide component, for example increasing the dosage of peroxide or using a more water-soluble peroxide than calcium peroxide, could also increase oxygen-delivery by polymer-peroxide composites. Thereby, a lower soluble peroxide might increase the release time.
In conclusion, biocompatible oxygen-releasing composite microspheres based on PTMC and CaO2 showed delayed skin necrosis in a random pattern
devascularized skin flap model in mice significantly for at least 10 days. These microspheres may support cell survival in otherwise ischemic circumstances and can aid in making the regeneration of tissue in ischemic environments more successful.
Funding
This research project was funded by Stichting voor Technische Wetenschappen, Utrecht, the Netherlands, grant number 10595.
Acknowledgements
Authors would like to thank Annemieke Smit-van Oosten, Bianca Meijeringh, Michel Weij and André Zandvoort, micro surgeons in the Animal Laboratory Groningen, for their excellent support. Authors would like to thank D.T.A. Ploeger, MSc. Ing., for assistance with immunostaining, Dr. S.M. van Putten for his assistance in the in vivo biocompatibility experiments and Marja Stiemsma-Slomp for her excellent laboratory support.
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