C57BL/6 (wild-type) mice and Tlr9^−/− mice (C57BL/6 background) were obtained from Japan SLC Inc. and Oriental BioService, Inc., respectively. Mice were maintained under a 12-h light/dark cycle, with a standard diet and water ad libitum. All experimental procedures conformed to the guidelines for animal experimentation of Tokushima University. The protocol was reviewed and approved by our institutional ethics committee under No. T29-96.

Unilateral hind-limb ischemia was induced in wild-type mice and Tlr9^−/− mice as described previously (17). Briefly, the animals were anesthetized by intraperitoneal injection of pentobarbital (50 mg/kg). The proximal and distal portions of the femoral artery and the distal portion of the saphenous artery were ligated. The arteries and all side branches were dissected free and excised. Blood flow in limbs was measured using a laser Doppler blood perfusion monitor (Moor LDPI) pre- and post-operatively and on days 7 and 14. Blood flow recovery in the ischemic limb was determined by the ratio of perfusion in the ischemic limb to the untreated limb.

Capillary density in the ischemic limb was measured as described previously (17). At 14 days after femoral artery ligation, mice were sacrificed by intraperitoneal injection of an overdose of pentobarbital. The whole limbs were fixed in methanol overnight, and the ischemic muscles were embedded in paraffin. Sections (5 μm) were de-paraffinized and incubated with an anti-CD31 antibody (clone MEC13.1, BD Pharmingen). Antibody distribution was detected with the avidinbiotin-complex technique and a Vector Red Alkaline Phosphatase substrate kit (Vector Laboratories). Nuclei were stained with hematoxylin. Six different fields from each section were randomly selected, and visible CD31-positive capillaries were quantitated with a FLOVEL Filing System (FLOVEL Company, Ltd.). Capillary density was expressed as the number of capillaries per field or per muscle fiber.

Accumulation of macrophages in ischemic limb was also examined by immunohistochemistry. Sections were incubated with anti-Mac3 antibody (BD Pharmingen). Development was performed by the combination of HRP-conjugated secondary antibody and ImmPACT DAB substrate (Vector Laboratories). Sections were counterstained with hematoxylin. Four different fields from each section were randomly selected, and Mac3-positive cells were quantitated with CellSens (OLYMPUS). Accumulation of macrophages was determined as the number of macrophages per field or per muscle fiber.

Thioglycollate-stimulated peritoneal macrophages were collected with cold PBS from age-matched female wild-type mice or Tlr9^−/− mice at the age of 8-12 weeks and cultured in DMEM containing 10% FBS at 37°C in a humidified incubator in 5% CO₂ and 95% air. At 24 h after plating, isolated peritoneal macrophages were used for experiments. Peritoneal macrophages were stimulated with ODN1826 or control-ODN1826 (GeneDesign, Inc.), synthetic oligonucleotides that contain unmethylated CpG, for indicated time. We collected plasma from wild-type mice at 3 days after femoral artery ligation or sham-operation. Wild-type or Tlr9^−/− macrophages were stimulated with these plasma for 4 h.

Human umbilical vein endothelial cells (HUVEC) were purchased from Life Technologies and grown in EGM-2 (Lonza) at 37°C in a humidified incubator in 5% CO₂ and 95% air. HUVEC (passages 4-6) were used for experiments.

HUVEC were plated in 96-well plates and grown to 40~50% confluence in EGM-2, and then stimulated with or without human recombinant TNF-α for 24 h in EBM-2 containing 1% FBS. Cell viability of HUVEC was determined by MTS assay using a CellTiter 96 AQueous One Solution Cell Proliferation Assay kit (Promega) according to the manufacturer's instructions. The percentage of absorbance was calculated against that of non-stimulated cells. Cell viability of HUVEC was also examined after treatment with conditioned medium (CM) of macrophages. Wild-type macrophages or Tlr9^−/− macrophages were pretreated with ODN1826 or control-ODN1826 (100 nM) under serum-starved conditions for 24 h, and then cells were cultured for another 24 h in a starvation medium without synthetic oligonucleotides. CM was collected after centrifugation and filtration through a 40-μm mesh to remove cell debris. Cell viability was determined by MTS assay after 72-h incubation.

cfDNA in plasma was extracted using a QIAamp DNA Mini Kit (Qiagen), according to the manufacturer's instructions. The concentrations of single-stranded DNA (ssDNA) and double-stranded DNA (dsDNA) in extracted cfDNA were measured using a QuantiFluor ssDNA System (Promega) and Quant-iT PicoGreen dsDNA Assay Kit (Life Technologies), respectively, according to the instructions.

RNA extracted from muscles and cells with an illustra RNAspin RNA Isolation Kit (GE Healthcare) was used for cDNA synthesis using a QuantiTect Reverse Transcription kit (Qiagen). Quantitative real-time RT-PCR (qPCR) was performed using Power SYBR Green PCR Master Mix (Applied Biosystems) and gene-specific primers on an Mx3000P (Agilent Technologies). Data are expressed in arbitrary units normalized by β-actin. The sequences of primers used in this study were as follows: TNF-α, sense 5′-accctcacactcagatcatcttc-3′ and antisense 5′-tggtggtttgctacgacgt-3′; F4/80, sense 5′-tgcatctagcaatggacagc-3′ and antisense 5′-gccttctggatcCATttgaa-3′; β-actin, sense 5′-CCTgagcgcaagtactctgtgt-3′ and antisense 5′-gctgatccacatctgctggaa-3′.

Protein lysates were isolated from tissues or cells using RIPA buffer (Wako Pure Chemical Industries, Ltd.) containing a protease inhibitor cocktail (Takara Bio Inc.) and phosphatase inhibitors (Roche LifeScience). Proteins were separated by SDS-PAGE and transferred to polyvinilidine difluoride membranes (Hybond-P; GE Healthcare). The membrane was blocked in 5% BSA for 1 h at room temperature, followed by incubation with a primary antibody against either total-IκBα (Cell Signaling Technology), TNF-α (abcam), α-Tubulin (MBL), or β-actin (Sigma) at 4°C overnight. After blots were washed in TBS containing 1% Tween-20, the membranes were incubated in horseradish peroxidase-conjugated secondary antibody (Chemicon) for 1 h. Expression of α-Tubulin or β-actin was used as an internal control to confirm equivalent total protein loading. Antibody distribution was visualized with ECL-plus reagent (GE Healthcare) using a luminescent image analyzer (LAS-1000, Fuji Film).

Data were expressed as mean ± SEM. Comparisons between two groups were made by unpaired Student's t test. Comparisons of multiple groups were made by one-way analysis of variance (ANOVA) followed by Tukey test. P < 0.05 was considered statistically significant.

To determine the participation of TLR9 in blood flow recovery of the ischemic limb, we examined the expression of TLR9 and circulating level of TLR9 ligand at 3 days after femoral artery ligation in wild-type mice. Results of qPCR demonstrated that ligation of the femoral artery markedly increased Tlr9 expression in the ischemic limb compared to the sham operated group (Figure 1A). Induction of hind-limb ischemia in wild-type mice also increased the plasma levels of dsDNA and ssDNA, internal ligands for TLR9, compared with the sham-operated group (Figures 1B,C).

Wild-type mice and Tlr9^−/− mice were subjected to femoral artery ligation, and serial blood flow measurements were performed by laser Doppler imaging. Tlr9^−/− mice showed better blood flow recovery than wild-type mice (Figure 2A). To clarify the role of TLR9 in the process of blood flow recovery, we examined capillary density in the ischemic limb by histological analysis. The results of CD31 staining showed that capillary density per square meter or per muscle fiber in the ischemic limb in Tlr9^−/− mice was higher than that in wild-type mice at 14 days after ligation (Figure 2B). We also found that Tlr9^−/− mice showed reduced infiltration of macrophages, one of the key players in the development of inflammation in the ischemic limb at 14 days after surgery, compared with wild-type mice (Figure 2C).

We examined the effect of TLR9 deficiency on the inflammatory process in the ischemic limb. The results of qPCR demonstrated that Tlr9^−/− mice had reduced expression of TNF-α and F4/80, a macrophage marker, in the ischemic limb at 3 days after surgery compared with wild-type mice (Figure 3A). The results of western blotting confirmed reduced expression of TNF-α in Tlr9^−/− mice at the protein level (Figure 3B). In addition, the results of western blotting using ischemic limbs showed that TLR9 deficiency significantly suppressed the degradation of IκBα compared with wild-type mice, indicating reduced activation of NF-κB signaling, one of the regulator TNF-α expression, in the ischemic limb in Tlr9^−/− mice (Figure 3C).

Inflammation associated with macrophage activation plays a pivotal role in blood flow recovery in the ischemic limb. Therefore, we examined the effect of TLR9 activation in macrophages. The results of in vitro experiments using peritoneal macrophages demonstrated that TLR9 activation by ODN1826, a synthetic oligonucleotide which activates TLR9, promoted the expression of TNF-α in wild-type macrophages, but not in Tlr9^−/− macrophages (Figure 4A). We further stimulated these macrophages with plasma obtained from mice which received femoral artery ligation or sham-operation. Wild-type macrophages increased TNF-α expression in the presence of plasma from mice with ischemic muscle compared with that from sham-operated mice, however, Tlr9^−/− macrophages did not show this response (Figure 4B). The ligation of TLR9 agonist to wild-type macrophages significantly activated NF-κB signaling as determined by the degradation of IκBα, however, this response was not observed in Tlr9^−/− macrophages (Figure 4C).

To investigate the effect of TLR9-mediated macrophage activation on endothelial cell, we treated HUVEC with the CM obtained from wild-type macrophages or Tlr9^−/− macrophages treated with ODN1826. CM obtained from wild-type macrophage treated with a TLR9 agonist significantly reduced the viability of HUVEC determined by MTS assay, although CM from Tlr9^−/− macrophage did not have any effect (Figure 5A). HUVEC treated with TNF-α also showed attenuated cell viability (Figure 5B). These results suggest that TNF-α, produced by macrophages via TLR9 activation, accelerated cell death of endothelial cells, leading to the suppression of blood flow recovery in ischemic limb.

The Department of Cardio-Diabetes Medicine, Tokushima University Graduate School, is supported in part by unrestricted research grants from Boehringer Ingelheim, Tanabe-Mitsubishi, Kowa, and Actelion. The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.