Abstract
This study aimed to clarify the phenotypical differentiation of recruited macrophages following subdermal implantation of an allogenous, acellular dermal matrix (aADM). In 20 male Wistar rats, one leg was randomly chosen for subcutaneous implantation of an aADM, while the other side received an autogenous dermis graft for control purposes. After 7 and 14 postoperative days, 10 animals were killed. Biopsies were obtained from the healing area and subjected to immunohistochemical staining (targets: pan macrophage marker CD68, M1 macrophage marker CD197, M2 macrophage marker CD163), histomorphometric analysis and reverse transcriptase polymerase chain reaction (targets: iNOS, arginase). No differences were detected in the total number of recruited macrophages between the groups. Allogenous ADMs significantly stimulated proinflammatory M1 differentiation, while autogenous dermis induced the regeneration promoting M2 phenotype. Proinflammatory M1 differentiation of macrophages might provide a potential explanation for profibrotic tissue deposition at the aADM interface following subcutaneous implantation, which has been observed previously.
Soft tissue augmentation with free, autogenous grafts is widely used in oral and maxillofacial surgery. The disadvantages of using autogenous grafts are mainly due to the harvesting procedure, which leads to a prolonged healing time at the donor site and therefore to increased patient morbidity . G riffin et al. investigated the morbidity following harvesting of free gingival grafts (FGG) or free subepithelial connective tissue grafts (SCTG) for gingival augmentation. They found postoperative pain, swelling and bleeding were the most important short term complications . Persistent numbness for several weeks after surgery was reported by D el P izzo et al. .
Anatomical and individual factors limit the amount of transplantable tissue that can be harvested safely . The quantity and quality of tissue that can be retrieved depends on the shape of the palatal vault, the patient’s sex and age. The location of the palatal vessels and nerves limits the amount of tissue available for grafting . An analysis of plaster casts revealed the average dimension of a FGG or SCTG to be 14.7 ± 2.9 mm × 31.7 ± 4.0 mm .
To overcome the problems of autogenous tissue transfer, alternative techniques employing allogenous and xenogenous materials have been reported. In a previous animal experimental study the authors demonstrated that allogenous, acellular, dermal matrices (aADMs) induce interfacial deposition of a dense connective tissue resulting in scarring and contraction at the implantation site .
It has been suggested that prolonged inflammation is the key factor triggering the fibrotic processes and that macrophages are the most important mediators of inflammatory reactions . Macrophages express CD68, a glycoprotein that binds to low density lipoprotein . As well as monocytes/macrophages, giant cells express the CD68 surface receptor . Polarized macrophages are referred to as M1 or M2 . M1, classically activated, proinflammatory macrophages are induced by factors such as interferon (INF)-γ or tumour necrosis factor (TNF), they express inducible nitric oxide synthetase (iNOS), interleukin (IL)-12, IL-23 and IL-10 and are involved in Th1 inflammatory responses . M1 macrophages are characterized by CD 197 (CCR 7; C-C chemokine receptor type 7), a G protein-coupled transmembrane receptor, which is activated in response to CCL19/MIP-3β or CCL21 and transduces chemotactic signals . This receptor is also expressed on mature dentritic cells, T- and NK-cells . In contrast, M2, alternatively activated macrophages are induced by factors such as IL-4 and IL-13, express arginase, IL-12, IL-23 and IL-10, are involved in polarized Th2 reactions and facilitate tissue repair and regeneration . M2 macrophages are characterized by CD163, a surface molecule belonging to the scavenger receptor superfamily , which mediates the up-take of haemoglobin–haptoglobin complexes. The molecule is exclusively expressed by monocytes/macrophages, but a soluble form can be released from the cell surface by proteolysis .
Taking into account fibrotic tissue deposition at the aADM interface on subcutaneous implantation and the data on macrophage phenotypes, this study tested the hypothesis whether predominant M1 differentiation of recruited macrophages takes place at the aADM implantation site.
Materials and methods
Animal model
20 male Wistar rats (Institute for Experimental Animals, University of Jena) with a mean age of 3 ± 1 months and a mean weight of 300–400 g were used. During the study, the animals were kept alone in Makrolon type III cages (Techniplast, Varese, Italy) at a temperature of 22 ± 0.5 °C, 55% humidity and a 12 h light/dark cycle. They received a pelleted standard rodent diet (N° 1320, Altromin, Lage, Germany) and fresh water ad libitum . The study protocol was approved by the Thuringian State Office for Food Safety and Consumer Care (Animal experiment application: 02-12/05).
The rats were anaesthetized with a mixture of ketamine (Ketavet ® , Pharmacia & Upjohn, Erlangen, Germany) and xylazine (Rompun ® , Bayer, Leverkusen, Germany) at a ratio of 2:1 (2.5 ml/kg body weight) by intraperitoneal injection. After making a 2 cm skin incision from the symphysis to the tibia in both legs, implants were placed on the gracilis muscle. One side was randomized to receive an aADM, which was prepared according to the dispase-triton method, as previously described , while the other received a free, autogenous dermis graft, harvested from the back of the rat. Both test materials were round in shape with a diameter of 1 cm. The thickness of both materials was not measured exactly, but both originated from rat back skin, and the thickness was estimated to be about 1 mm . Ten animals were killed after 7 and 14 postoperative days. Biopsies were retrieved from the healing area. Each biopsy was bisected vertically. One half of each biopsy was fixed in 4% formalin and the other half was stored in RNAlater (QIAGEN, Hilden, Germany) for reverse transcriptase polymerase chain reaction (RT-PCR) analysis.
Haematoxylin-eosin (HE) staining
One half of each biopsy was fixed in 4% formalin and embedded in paraffin (Histokinette, Leica, Nussloch, Germany). Paraffin-embedded specimens were cut vertically into 3 μm sections using the microtome (RM2145, Leica, Nussloch, Germany). Three consecutive sections per specimen were placed on one slide. The slides were subjected to standard HE staining according to the method described by R omeis et al. . In brief, slides were incubated in haemalaun according to Mayer for 25 min (1 mg/ml haematoxylin, 200 μg/ml sodium iodide, 50 mg/ml potassium alaun, 50 mg/ml chloral hydrate, 1 mg/ml citric acid; Merck, Darmstadt, Germany), rinsed with distilled water for 5–10 min, incubated in eosin (1 mg/ml; Merck, Darmstadt, Germany) for 10 min and rinsed with distilled water. HE-stained sections were examined qualitatively under a bright field microscope (Axioskop, Zeiss, Oberkochen, Germany) at magnifications of 50× and 100×.
Immunohistochemical staining
Biopsy specimens were subjected to immunohistochemical staining for the pan macrophage marker CD68, the M1 macrophage marker CD197 and the M2 macrophage marker CD163 using the avidin–biotin–peroxidase complex (ABC-POX) method as described previously . Endogenous peroxidase activity was blocked by incubation in 3% H 2 O 2 for 20 min at room temperature. The specific epitopes were demasked with citrate buffer (Dako, Carpinteria, CA, USA) at 90 °C for 25 min. Preparations were incubated for 30 min in blocking solution (500 mg skimmed milk powder, TBS, 0.1% Tween 20; Merck, Germany).
CD68 was marked by incubation with a monoclonal, unconjugated mouse-IgG 1 (AbD Serotec, Oxford, UK; dilution: 1:50; overnight; 4 °C). CD197 was stained with a polyclonal rabbit-IgG antibody (Cell Applications, San Diego, CA, USA; dilution: 1:100; overnight; 4 °C) and CD163 was detected using a monoclonal mouse-IgG 1 (AbD Serotec; dilution: 1:50; overnight; 4 °C).
Rabbit-anti-mouse IgG (Dako; dilution: 1:250; 30 min; room temperature) was used as a secondary antibody for CD68 and CD163 staining, while a polyclonal goat-anti-rabbit antibody (Dako; dilution: 1:80; 30 min; room temperature) was used for CD197 staining. For chromogenic development avidin–biotin/horseradish peroxidase complex (ABC/HRP complex, Dako; 30 min, room temperature) was used and AEC (0.02% 3-amino-9-ethylcarbazole in 50 mM acetate buffer pH 5; 5.5% dimethylformamide, Dako) served as a substrate. Cell nuclei were counterstained using haematoxylin (Dako). The antibodies were specific. From each tissue sample, three consecutive sections were incubated on glass slides, with one negative control (omitting primary antibody).
Histomorphometric analysis
Slides were examined qualitatively under a bright field microscope (Axioskop, Zeiss, Oberkochen, Germany) with 100–400× magnification for density and localization of CD68-, CD197- and CD163-positive cells in the healing area, which was defined as the region between the gracilis muscle at the bottom and the epithelium at the top. In case of epithelial destruction by the cutting process, as present in the aADM group, the apical connective tissue surface was defined as the border of the healing area. This was determined descriptively and compared with the control checks.
The total number of CD68-, CD197- and CD163-positive cells per visual field was counted and the ratios of CD197- and CD68-positive cells as well as CD163- and CD68-positive cells were determined. Three, randomly chosen, high power fields (HPF) per section for each sample, animal, day, and group were digitized with 400× magnification using a charge-coupled device camera (Axiocam, Zeiss, Oberkochen, Germany) and the program Axiovision Rel. 4.5 (Zeiss, Oberkochen, Germany). The evaluation was carried out independently by three examiners. The criteria for positively stained cells were the existence of a clear cell structure with a nucleus and clear specific staining of the cytoplasm. Staining of the extracellular matrix was not evaluated.
RNA-extraction and qRT-PCR
Total RNA was extracted from the samples using the RNeasy Mini Kit (QIAGEN, Hilden, Germany). RNA concentrations were determined spectrophotometrically (NanoDrop ® , Wilmington, DE, USA). In order to confirm M1 or M2 macrophage phenotype quantitative, real-time one-step RT-PCR was performed for the genes iNOS and arginase (ARG) using the Mastercycler ® ep realplex system (Eppendorf, Hamburg, Germany) and the QuantiTect Primer Assay kit (Qiagen, Hilden, Germany). Briefly, 1 ng/μl RNA, 10 μl 2× QuantiTect SYBR Green RT-PCR Master Mix, 10 μl 10× QuantiTect Primer Assay primers for iNOS (QT01323189; Qiagen) or ARG (QT00177611; Qiagen) or GAPDH (QT00199633; Qiagen) and 0.2 μl QuantiTect RT Mix were mixed and DNase-/RNase-free water was added up to a total volume of 20 μl in 96-well plates. The PCR reaction cycles comprised 30 min at 50 °C and 15 min at 95 °C, followed by 40 cycles of 15 s at 94 °C, 20 s at 55 °C and 20 s 72 °C. Data were analysed by the ΔΔ-Ct method . In brief, concentration in time (Ct) values of the genes of interest (iNOS, ARG) were corrected for Ct values from the housekeeping gene GAPDH, resulting in a ΔCt value (ΔCt = Ct target gene − Ct housekeeping gene (GAPDH) ). ΔΔ-Ct was calculated by subtracting the ΔCt of the control group (autogenous dermis graft) from the ΔCt of the experimental group (aADM), (ΔΔCt = ΔCt experimental group − ΔCt control group ). Relative mRNA levels were calculated by 2 −ΔΔCt for each target gene. All reactions were performed in duplicate for each sample.
Statistics
Mean value ± SD were calculated. Graphic description was carried out using the bar-plot. Comparison of the total number of CD68-positive cells per HPF as well as the percentage of CD197- and CD163-positive macrophages between both sides at different time points was made using unpaired t-tests. Two sided p -values of p ± 0.05 were considered significant. All calculations were made using SPSS V.15.0 for Windows.
Results
Qualitative results
HE-stained sections at 50× and 100× magnification were chosen for cross-sectional analysis of the healing area, which was defined as the region between the gracilis muscle at the bottom and the epithelium at the top. Following transplantation of autogenous dermis into the groin region, the graft was not distinguishable from the surrounding tissue at day 7 or 14 postoperatively ( Fig. 1 A ) and a fibrotic connective tissue layer was formed around the aADMs ( Fig. 1 B).
To investigate monocytic infiltration in the healing area, sections stained with the pan macrophage-marker CD68 were analysed at 400× magnification. At postoperative day 7, the host response to the grafting procedures was characterized by a dense monocytic infiltration, mainly localized in the perivascular area. Following grafting of an aADM, the monocytic infiltration seemed to be more pronounced compared with autogenous dermis grafting ( Fig. 2 A ). Fourteen days postoperatively the distribution of the cells changed from a perivascular to an interstitial pattern. The staining seemed more intense following aADM transplantation ( Fig. 2 B).
To allow for further discrimination of the phenotypical polarization of the recruited monocytes/macrophages, CD197 (proinflammatory M1 macrophages) and CD163 (regenerative/reparative macrophages) stained sections were analysed at 400× magnification. At day 7 postoperatively only a few macrophages exhibited polarization. Following aADM grafting, CD197 seemed more intense compared with autogenous dermis grafting ( Fig. 2 C). An inverse situation was observable for CD163 ( Fig. 2 E). At day 14 postoperatively, significant polarization was present. CD197 was highly expressed following aADM grafting ( Fig. 2 D) while CD163 was more intense following autogenous dermis grafting ( Fig. 2 F).
Quantitative analysis
Quantification of the total number of macrophages by counting all CD68-positive cells per HPF revealed no significant differences between autogenous dermis grafts and aADMs during the entire investigation period. Following transplantation of autogenous dermis grafts, 143 ± 31 macrophages/HPF were detectable 7 days postoperatively and 123 ± 31 macrophages/HPF were counted 14 days postoperatively in the healing area. Allogenous ADM grafted sides showed mean macrophage numbers of 163 ± 35/HPF at day 7 postoperatively and 117 ± 40/HPF at day 14 postoperatively ( Fig. 3 A ).
