OtherBASIC SCIENCE INVESTIGATIONS

Inflammatory Cytokines and Hypoxia Contribute to 18F-FDG Uptake by Cells Involved in Pannus Formation in Rheumatoid Arthritis

Tamiko Matsui, Norihito Nakata, Shigenori Nagai, Akira Nakatani, Miwako Takahashi, Toshimitsu Momose, Kuni Ohtomo and Shigeo Koyasu
Journal of Nuclear Medicine June 2009, 50 (6) 920-926; DOI: https://doi.org/10.2967/jnumed.108.060103

Abstract

Assessment of the activity of rheumatoid arthritis (RA) is important for the prediction of future articular destruction. 18F-FDG PET is known to represent the metabolic activity of inflammatory disease, which correlates with the pannus volume measured by MRI or ultrasonography. To evaluate the correlation between 18F-FDG accumulation and RA pathology, we assessed 18F-FDG accumulation in vivo using collagen-induced arthritis (CIA) animal models and 3H-FDG uptake in vitro using various cells involved in arthritis. Methods: 18F-FDG PET images of rats with CIA were acquired on days 10, 14, and 17 after arthritis induction. The specimens were subsequently subjected to macroautoradiography, and the 18F-FDG accumulation was compared with the histologic findings. 3H-FDG uptake in vitro in inflammatory cells (neutrophils, macrophages, T cells, and fibroblasts) was measured to evaluate the contributions of these cells to 18F-FDG accumulation. In addition, the influence on 3H-FDG uptake of inflammatory factors, such as cytokines (tumor necrosis factor α [TNFα], interleukin 1 [IL-1], and IL-6), and hypoxia was examined. Results: 18F-FDG PET depicted swollen joints, and 18F-FDG accumulation increased with the progression of arthritis. Histologically, a higher level of 18F-FDG accumulation correlated with the pannus rather than the infiltration of inflammatory cells around the joints. In the in vitro 3H-FDG uptake assay, fibroblasts showed the highest 3H-FDG uptake, followed by neutrophils. Although only a small amount of 3H-FDG was incorporated by resting macrophages, a dramatic increase in 3H-FDG uptake in both fibroblasts and macrophages was observed when these cells were exposed to inflammatory cytokines, such as TNFα and IL-1, and hypoxia. Although neutrophils showed relatively high 3H-FDG uptake without activation, no increase in 3H-FDG uptake was observed in response to inflammatory cytokines. 3H-FDG uptake by T cells was much lower than that by other cells. Thus, fibroblasts and activated macrophages contribute to a high level of 18F-FDG accumulation in the pannus, and hypoxia as well as cytokine stimulation significantly increases 18F-FDG uptake by these cells. Conclusion: 18F-FDG accumulation in RA reflects proliferating pannus and inflammatory activity enhanced by inflammatory cytokines and hypoxia. 18F-FDG PET should be effective for quantifying the inflammatory activity of RA.

Rheumatoid arthritis (RA) is an autoimmune disorder of unknown etiology and is characterized by systematic, symmetric, and erosive synovitis (1). RA synovitis shows massive leukocyte infiltration, proliferative synovial membranes, and neovascularization, which give rise to a synovial hypertrophy called a pannus. Such pannus formation is directly responsible for cartilage and bone destruction (2,3).

It recently became clear that the erosive changes of bone appear very early in RA (4), that the treatment of RA should start as early as possible, and that RA must be controlled adequately to reduce the occurrence of irreversible joint damage (5). Therefore, the early identification of pathologic synovitis and the appropriate monitoring of RA disease activity are important for the control of RA. However, current assessments of RA activity are not sufficient for appropriate monitoring. The conventional estimation of clinical features, such as joint tenderness and swelling, are relatively subjective, and laboratory examinations, such as the erythrocyte sedimentation rate (ESR) and serum C-reactive protein level, show just the mean value of the systemic inflammatory state. Radiography is unable to directly evaluate synovial inflammation before bone destruction (6).

For overcoming these problems, some imaging modalities are capable of quantifying the activity of synovitis in each joint; these include MRI, high-resolution ultrasonography (US), and PET (7–9). PET imaging with 18F-FDG, which is a glucose analog that reflects glucose metabolism, has been widely used in clinical oncology (10). Lately, it has been shown that 18F-FDG PET can be applied to the evaluation of inflammatory diseases. For RA, it has been reported that 18F-FDG uptake represents the metabolic activity of synovitis (8). Furthermore, correlations among the 18F-FDG accumulation in synovitis, the volume of the pannus evaluated by MRI or US, and the therapeutic response have been shown (7,9,11). In addition, an advantage of 18F-FDG PET is that whole-body 18F-FDG PET can evaluate the activity of synovitis in all joints in a single examination (12).

However, the mechanisms of how 18F-FDG accumulates in the inflammatory region, what types of cells play a major role in incorporating 18F-FDG, and how inflammatory reactions affect the accumulation of 18F-FDG are largely unknown. In this study, we examined the mechanisms of 18F-FDG accumulation by using a murine collagen-induced arthritis (CIA) model in vivo and 3H-FDG uptake by various cell types in vitro. We also examined the effects of inflammatory cytokines and hypoxia, both of which are known to affect the characteristics of various cells in inflammation. Our results show that both macrophages and fibroblasts play a major role in the uptake of 3H-FDG and that inflammatory cytokines and hypoxia greatly enhance 3H-FDG uptake in vitro, which likely reflects 18F-FDG uptake in vivo.

MATERIALS AND METHODS

Preparation of CIA Model

Six-week-old female Lewis rats were purchased from Japan SLC and housed under constant environmental conditions with 12-h light–dark cycles. Food and water were provided ad libitum. Arthritis was induced according to the protocol recommended by Chondrex Inc., the supplier of bovine type II collagen. In brief, collagen (2 mg/mL) was mixed with an equal volume of N-acetylmuramyl-l-alanyl-d-isoglutamine hydrate (MDP; 8 mg/mL; Sigma). The collagen–MDP solution was emulsified with an equal volume of incomplete Freund adjuvant (Sigma) by stirring with an electric homogenizer. Rats were injected intradermally in the base of the tail with 0.3 mL of the emulsion of collagen–MDP–incomplete Freund adjuvant. The severity of arthritis in each paw was evaluated according to the following scale: 0 = no swelling or redness (normal paw); 1 = swelling or redness in digits; 2 = focal swelling or redness in digits and paw; 3 = swelling in entire paw; and 4 = severe swelling in entire paw. Animals that did not receive collagen injections were used as controls.

18F-FDG PET

All rats were fasted for 16–20 h before the experiments but had free access to water. After thiopental sodium anesthesia (40 mg/kg intraperitoneally), rats were injected in the tail vein with 37 MBq (1.0 mCi) of 18F-FDG (Nihon Medi-physics). Forty minutes after 18F-FDG injection, emission scans were performed for 10 min with a small-animal PET scanner (eXplore Vista DR; GE Healthcare). The PET images were reconstructed by use of a 2-dimensional ordered-subset expectation maximization algorithm with 32 subsets, 2 iterations, random and scatter corrections, and no attenuation correction.

Macroautoradiography and Histologic Analysis

Rats were sacrificed 1 h after the 18F-FDG injection (i.e., 10 min after the PET scan). Frozen sections of joints were prepared by use of the film method developed by Kawamoto et al. (13). In brief, hind paws were frozen in liquid nitrogen and embedded in cryoembedding medium (Finetec). Adhesive film (Cryofilm; Finetec) was attached to the exposed surface of frozen samples, and 10-μm-thick sections were cut with a cryostat (CM3050S-10; Leica Instruments) and mounted on glass slides. For macroautoradiography, the slides were exposed to imaging plates (BAS-SR 2040; Fuji Film) overnight. The imaging plates were then scanned with an imaging analyzer (BAS-2500; Fuji Film). Sections were then stained with hematoxylin and eosin (HE) for histologic evaluation. Immunohistochemical staining was performed on sections of day 17 samples. Contiguous sections were incubated with an anti–rat CD68 antibody (BMA Biomedicals) and an anti–rat granulocyte antibody (BD Biosciences Pharmingen) overnight at 4°C. Bound antibodies were detected with an LSAB2 Streptavidin-HRP (Dako) and 3,3′-diaminobenzidine tetrahydrochloride.

Preparation of Cells

Six-week-old male C57BL/6 mice were purchased from Japan SLC. Bone marrow cells collected from femurs were suspended in serum-free RPMI 1640 medium, and red blood cells were removed by use of the Lymphosepar (Immuno-Biologic Laboratories) density gradient method according to the manufacturer's recommendations. Mononuclear cells were cultured with 10 nM mouse macrophage–colony-stimulating factor (M-CSF; R&D) in RPMI 1640 culture medium supplemented with 10% fetal calf serum, 55 μM 2-mercaptoethanol, penicillin at 100 U/mL, streptomycin at 100 μg/mL, nonessential amino acids (Invitrogen), 1 mM sodium pyruvate, and 10 mM N-2-hydroxyethylpiperazine-N′-(2-ethanesulfonic acid) (pH 7.4) at 37°C in a 5% CO2 humidified incubator. After 24 h of cultivation, nonadherent cells were collected and cultured for an additional 7 d in culture medium containing 40 nM M-CSF. Adherent cells were collected as bone marrow–derived macrophages and used for experiments.

Neutrophils were obtained from bone marrow cells by Histopaque (Sigma) density gradient centrifugation. Histopaque 1.083 was added on top of Histopaque 1.119, and the cell suspension was placed on the upper layer and centrifuged at 700g for 30 min at room temperature. This method routinely yielded nonprimed and viable neutrophils with greater than 85% purity.

Fibroblasts were isolated from dermal tissues. The back skin of a C57BL/6 mouse was washed with 70% ethanol, and the epidermis and dermis were separated physically after digestion with 0.3% (w/v) trypsin for 60 min at 37°C. The dermal samples were cut into small (2- to 3-mm) square pieces, and a few skin pieces were incubated under glass cover slips in a 6-well plate with RPMI 1640 culture medium. When fibroblast outgrowth became confluent, the cover slips and skin pieces were removed, and fibroblasts were collected with 0.25% trypsin–ethylenediaminetetraacetic acid at room temperature.

T cells were purified from splenocytes with anti–mouse CD90-conjugated magnetic beads (Miltenyi Biotec) and Auto MACS (Miltenyi Biotec) according to the manufacturer's instructions. T cells were activated with concanavalin A (Con A) at 4 μg/mL and cultured for 7 d to prepare preactivated T cells in a resting state.

Measurement of 3H-FDG Uptake

3H-FDG uptake in vitro was examined with 3.7 kBq of 5,6-3H-FDG (American Radiolabeled Chemicals) in assay medium (serum- and glucose-free RPMI 1640 medium, pH 7.4) for 1 h at 37°C. The effects of inflammatory cytokines on 3H-FDG uptake were evaluated with the following cytokines: tumor necrosis factor α (TNFα; 100 ng/mL; R&D), interleukin 1β (IL-1β; 1,000 U/mL; R&D), and IL-6 (100 ng/mL; PeproTech). We also used phorbol myristate acetate (PMA; 10 ng/mL; Sigma), a known activator of neutrophils. Macrophages (2 × 105 cells/300 μL/2 cm2) and fibroblasts (5 × 104 cells/300 μL/2 cm2) were incubated in culture medium with or without cytokines for 24 h, and then the culture media were replaced with 500 μL of assay medium. Neutrophils (5 × 105 cells/500 μL/1.5-mL tube) were incubated in assay medium with or without stimulation.

After incubation with 3H-FDG, uptake was terminated with 3 washes of cold phosphate-buffered saline. The cells were then solubilized with 0.9 mL of 1N NaOH and neutralized with 0.9 mL of 1N HCl. Radioactivity was determined by use of a liquid scintillation counter (Tri-CARB 3100; Perkin-Elmer). The uptake of 3H-FDG was calculated from the ratio of 3H-FDG associated with the cell pellet to the initial dose (percentage injected dose [%ID]) and was normalized to living cell numbers (%ID/105 cells).

For hypoxia treatment, the cells were cultured in a humidified multigas incubator (APM-30DR; Astec) with 1% O2 (hypoxia) and 5% CO2 at 37°C for 24 h. Assay medium was preequilibrated overnight before use. Cells cultured in normoxia (20.9% O2) were used as controls.

Statistics

All values are expressed as the mean ± SD of multiple independent experiments. The data were evaluated statistically with the Mann–Whitney U test. Differences were considered significant when the P value was less than 0.05.

RESULTS

18F-FDG Accumulation in CIA Model In Vivo

To evaluate how 18F-FDG PET depicts arthritis during the progress of the disease, PET and macroautoradiography of rats with CIA were performed at various days after the induction of arthritis. Rats developed arthritis at about day 14 after collagen injection and developed severe swelling of the joints by day 17 (Supplemental Fig. 1) (supplemental materials are available online only at http://jnm.snmjournals.org). We examined 18F-FDG uptake in the joints by 18F-FDG PET, and representative images are shown in the top row of Figure 1. The PET image of rats with CIA on day 10 was indistinguishable from that of control rats and revealed no marked 18F-FDG accumulation in their joints. On day 14, when the disease symptoms were readily recognized, the swollen joints were clearly visualized by PET. The accumulation of 18F-FDG in the tarsal joints was further increased by day 17, the later stage of the disease. These results indicated that 18F-FDG PET is able to depict inflamed joints and that 18F-FDG accumulation increases with the progression of arthritis.

FIGURE 1. 
FIGURE 1. 

18F-FDG PET images of CIA animal model. PET images acquired 40 min after injection of 37 MBq of 18F-FDG into rats with CIA (top row), macroautoradiograms (ARG) of sagittal sections (middle row), and HE staining of tarsometatarsal joints (bottom row) are shown. Images represent control rat and rats with CIA 10, 14, and 17 d after collagen injection. Bar = 200 μm.

We performed macroautoradiography of the joints after obtaining PET images (Fig. 1, middle row). Specimens on day 14 showed the focal accumulation of 18F-FDG around the joints, and those on day 17 showed a high level of accumulation of 18F-FDG at the advanced arthritis region (Fig. 1, middle row). Histologic analyses (Fig. 1, bottom row) revealed no significant changes in the joints 10 d after collagen injection. Rats with CIA on day 14 showed synovial cell hyperplasia with edema and interstitial infiltration of inflammatory cells—mainly neutrophils and macrophages. On day 17, panni consisting of mixed cellular patterns of macrophages and fibroblasts were observed, and bone destruction by mature osteoclasts was often observed (Fig. 1, bottom row). Similar results were obtained with the CIA mouse model (Supplemental Fig. 2).

18F-FDG Accumulation in Regions of Pannus and Bone Destruction

When we compared the histologic findings with 18F-FDG accumulation in detail, the areas with a high level of 18F-FDG accumulation (Fig. 2A, arrow) corresponded to the regions of pannus and bone destruction (Fig. 2B). In contrast, the areas with a moderate level of 18F-FDG accumulation (Fig. 2A, arrowhead) corresponded to synovial cell hyperplasia and edematous inflammation (Fig. 2E). These observations indicated that the distribution of 18F-FDG corresponds to inflammatory regions, such as those with pannus formation and the infiltration of inflammatory cells. A high level of 18F-FDG accumulation especially tends to correlate with severe pannus formation and considerable bone destruction.

FIGURE 2. 
FIGURE 2. 

Immunohistochemical analyses of joints affected by CIA. (A) Macroautoradiogram of rat with CIA 17 d after collagen injection. (B and E) HE staining in region with high level of 18F-FDG accumulation (B, arrow in A) and region with moderate level of 18F-FDG accumulation (E, arrowhead in A). Asterisk in B shows bone destruction with pannus. Arrows in E show synovial cell hyperplasia. Bar = 500 μm. (C, D, F, and G) Representative immunohistochemical staining of serial sections for CD68 (C and F) and granulocytes (D and G). Data are from region with high level of 18F-FDG accumulation (C and D; enlarged view of square in B) and region with moderate level of 18F-FDG accumulation (F and G; enlarged view of square in E). Bar = 50 μm.

Immunohistochemical staining was performed to examine the contributions of inflammatory cells to 18F-FDG accumulation. Neutrophils and macrophages were readily observed to similar degrees in areas with both high and moderate levels of 18F-FDG accumulation (Figs. 2C, 2D, 2F, and 2G), suggesting that the number of inflammatory cells may not be the major cause of localized 18F-FDG accumulation in tissues with arthritis. It is possible that the difference in 18F-FDG accumulation is attributable to the presence of fibroblasts as a constituent of the pannus. Alternatively, the increase in glucose metabolism observed with inflammatory cells may occur in a distinct microenvironment. For example, stimulation of arthritis with local cytokines or hypoxia may cause a high level of accumulation of 18F-FDG.

3H-FDG Uptake by Various Cells In Vitro

To examine the factors affecting 18F-FDG uptake by various cell types involved in the pathogenesis of RA (macrophages, neutrophils, fibroblasts, and T cells) in vivo, we performed a series of experiments with 3H-FDG to evaluate the ability of various cells to incorporate 18F-FDG and the effects of various inflammatory cytokines (TNFα, IL-1, and IL-6) in vitro. Preincubation of macrophages with TNFα resulted in a large increase in 3H-FDG uptake (TNFα, 0.56 ± 0.25 %ID/105 cells; control, 0.11 ± 0.05 %ID/105 cells; P < 0.05), whereas IL-1 and IL-6 had little effect (Fig. 3A). For fibroblasts, stimulation by TNFα or IL-1 significantly increased 3H-FDG uptake (TNFα, 2.21 ± 0.24 %ID/105 cells; IL-1, 1.71 ± 0.21 %ID/105 cells; control, 0.93 ± 0.16 %ID/105 cells; P < 0.05) (Fig. 3C). In contrast, the incubation of neutrophils (5 × 105 cells per tube) with TNFα, IL-1, or IL-6 for 1 h resulted in no significant increase in 3H-FDG uptake, whereas a significant increase in 3H-FDG uptake was observed when neutrophils were stimulated with the phorbol ester PMA (PMA, 0.82 ± 0.35 %ID/105 cells; control, 0.49 ± 0.14 %ID/105 cells; P < 0.05) (Fig. 3B). These results indicated that 3H-FDG uptake by neutrophils is not significantly affected by these inflammatory cytokines. 3H-FDG uptake by both naive T cells and T cells preactivated with Con A for 7 d was very low compared with that observed with other inflammatory cells (Fig. 3D and data not shown).

FIGURE 3. 
FIGURE 3. 

3H-FDG uptake in vitro and effects of inflammatory cytokines. (A–C) Effects of inflammatory cytokines on 3H-FDG uptake in macrophages (A), neutrophils (B), and fibroblasts (C). Macrophages and fibroblasts were cultured with TNFα (100 ng/mL), IL-1 (1,000 U/mL), or IL-6 (100 ng/mL) for 24 h and then incubated with 3H-FDG for 1 h. Neutrophils were incubated with 3H-FDG for 1 h along with TNFα (100 ng/mL), IL-1 (1,000 U/mL), IL-6 (100 ng/mL), or PMA (10 ng/mL). 3H-FDG uptake was normalized to numbers of living cells. Data represent mean ± SD of percentage of uptake of initial dose per 105 cells in 3 separate experiments. (D) Comparison of 3H-FDG uptake in different types of cells. *P < 0.05 for comparison with control (n = 3 for each group), as determined by Mann–Whitney U test.

3H-FDG uptake was highest for fibroblasts (0.93 ± 0.16 %ID/105 cells), followed by neutrophils (0.49 ± 0.14 %ID/105 cells), macrophages (0.11 ± 0.05 %ID/105 cells), and T cells (0.03 ± 0.0 %ID/105 cells) (Fig. 3D). These results indicated that fibroblasts significantly contribute to 18F-FDG uptake and that inflammatory cytokines, such as TNFα, significantly enhance 18F-FDG uptake by fibroblasts and macrophages.

Hypoxia Enhances 3H-FDG Uptake

Because it is known that inflammatory areas, including those in RA, are hypoxic compared with noninflammatory areas (14–16), we examined the effect of hypoxia on 3H-FDG uptake by various cells (Fig. 4). Under hypoxic conditions, 3H-FDG uptake by macrophages (hypoxia, 1.2 ± 0.42 %ID/105 cells; normoxia, 0.29 ± 0.31 %ID/105 cells; P < 0.05) (Fig. 4A) and fibroblasts (hypoxia, 10.03 ± 2.27 %ID/105 cells; normoxia, 1.62 ± 0.38 %ID/105 cells; P < 0.05) (Fig. 4B) was significantly increased. 3H-FDG uptake was further enhanced by TNFα stimulation under hypoxic conditions for macrophages (hypoxia, 3.53 ± 1.43 %ID/105 cells; normoxia, 1.20 ± 0.42 %ID/105 cells; P < 0.05) and fibroblasts (hypoxia, 16.65 ± 4.17 %ID/105 cells; normoxia, 10.03 ± 2.27 %ID/105 cells; P < 0.05) (Fig. 4). Interestingly, 3H-FDG uptake by neutrophils was not enhanced under hypoxic conditions (data not shown). These results indicated that hypoxia as well as cytokine stimulation significantly increases 3H-FDG uptake by macrophages and fibroblasts.

FIGURE 4. 
FIGURE 4. 

Effects of hypoxia on 3H-FDG uptake in macrophages (A) and fibroblasts (B). Macrophages and fibroblasts were incubated in absence (open bars) or presence (filled bars) of TNFα (100 ng/mL) in normoxia (20.9% O2) or hypoxia (1% O2) for 24 h. Cells were then incubated with 3H-FDG for 1 h. Data represent mean ± SD of 3 separate experiments. *P < 0.05 in comparison with nonstimulated control (n = 3); †P < 0.05 in comparison with nonstimulated sample in normoxia (n = 3). P values were determined by Mann–Whitney U test.

DISCUSSION

It was previously shown that 18F-FDG can be used to assess the activity of rheumatoid synovitis and that the standardized uptake value is closely correlated with the volume of the enhancing pannus or the synovial thickness evaluated by MRI and US (7,9,11). However, it has not been clear what pathologic condition of RA 18F-FDG reflects. In the present study, animal models of arthritis was used to show that 18F-FDG accumulation in arthritis correlated with the active pannus. Fibroblasts showed the highest level of 3H-FDG uptake among the various cell types forming the pannus. Our results also demonstrated that 3H-FDG uptake was increased in the presence of proinflammatory cytokines. It has been shown that TNFα and IL-1 induce the cell cycle progression of fibroblasts (17,18). In fact, synovial fibroblasts in joints affected by RA showed increased proliferative activity and increased glycolysis compared with those in normal joints (19,20). Fibroblast proliferation may be one of the dominant causes of 18F-FDG accumulation in vivo. Thus, it is suggested that fibroblasts play a key role in 18F-FDG accumulation in arthritis.

Previous studies suggested that neutrophils and macrophages contribute to 18F-FDG accumulation mainly in inflammatory diseases (17,21). Indeed, although only a small amount of 3H-FDG was incorporated by resting macrophages, a dramatic increase in 3H-FDG uptake was observed on stimulation with TNFα. Activated macrophages thus contribute to 18F-FDG accumulation in RA. In contrast, neutrophils showed a relatively high level of 3H-FDG uptake without activation, but inflammatory cytokines had little influence on the 3H-FDG uptake. These results indicate that the contribution of neutrophils to 18F-FDG accumulation depends simply on the amount of cell infiltration and does not reflect the activity of inflammatory cytokines.

On the basis of our results, it is likely that abundant cytokine production in RA is one of the characteristic conditions that affect 18F-FDG uptake. TNFα, IL-1, and IL-6 are known to be the major cytokines involved in the process of RA. These cytokines are produced by synoviocytes and mononuclear cells and activate inflammatory cells as well as fibroblasts (18,22). The energy to support the inflammatory activities enhanced by these cytokines is supplied primarily by glucose metabolism, as the depletion of glucose from the medium easily gives rise to a severe depression of IL-1 production from stimulated macrophages (23).

In addition to the production of inflammatory cytokines, another characteristic of RA is hypoxia. The dysfunction of the neovascular network in RA fails to maintain tissue oxygen homeostasis, and intense inflammation induces marked hypoxia in joints affected by RA (14–16). We showed here that hypoxia is another condition that enhances the uptake of 18F-FDG by both macrophages and fibroblasts. Hypoxia changes the energy supply of cells into anaerobic glycolysis. According to studies of glucose use by macrophages, differences in O2 tension lead to alterations in the rate of oxidative phosphorylation and glycolysis, and hypoxia upregulates glucose uptake and lactate production (24,25). Furthermore, it is known that macrophages regulate many factors associated not only with survival, such as the glucose transporter and vascular endothelial growth factor, but also inflammation, such as TNFα and IL-1, under hypoxic conditions (26). Cellular activation induced by hypoxia is a critical factor that complicates synovitis and increases 18F-FDG accumulation in RA. In contrast, because neutrophils are thought to provide glycolytic substrates from intracellular glycogen in the presence of low glucose levels, it is possible that 3H-FDG uptake in neutrophils is not affected by activation under experimental conditions (27,28).

For human RA, fewer neutrophils but large numbers of T cells and B cells infiltrate the pannus; this pattern is different from the observations in the CIA animal model. The CIA animal model shows a pathologically acute inflammatory condition associated with marked neutrophil invasion; this pattern is rare in human RA. Because of the involvement of fewer neutrophils in human RA, it is believed that 18F-FDG PET can depict the proliferating pannus more specifically. In contrast, to fibroblasts the level of 3H-FDG uptake by T cells preactivated with Con A was much lower than that by other cells, suggesting that T cells do not play a major role in 18F-FDG accumulation in synovitis. Furthermore, B cells observed in rheumatoid synovitis often form germinal centers in the region of rheumatoid synovitis and proliferate in situ (29). Because B cells accumulate 18F-FDG under proliferative conditions (30), the involvement of ectopic germinal centers of such proliferating B cells would contribute to 18F-FDG accumulation in rheumatoid synovitis in human RA. In addition, it is known that the extent of ectopic germinal centers in B cells varies among individuals; thus, 18F-FDG accumulation in human RA may be affected by the extent of ectopic germinal centers in B cells. These issues remain to be investigated in future studies.

Our data support the results of previous clinical studies showing that the standardized uptake value correlates with the volume of the pannus evaluated by MRI and US. Our data also show that 3H-FDG uptake varies depending on the environment of the cells forming the pannus. These results suggest that 18F-FDG accumulation in RA reflects the inflammatory environment that results from the disease activity as well as the volume of the pannus. In clinical examinations of RA, because the activated pannus directly leads to the destruction of articular bone and cartilage, 18F-FDG accumulation in RA may provide important biologic information about future joint destruction; in addition, 18F-FDG PET may have advantages over MRI or US for evaluation of the disease activity of RA.

CONCLUSION

We evaluated the correlation between 18F-FDG accumulation in arthritis and pathologic findings by using a CIA animal model; we found a higher level of 18F-FDG accumulation in the pannus. The in vitro 3H-FDG uptake assay with various cells involved in the pannus indicated that fibroblasts predominantly contributed to 18F-FDG accumulation. Additionally, inflammatory conditions, such as inflammatory cytokine production and hypoxia, altered cellular activity or metabolism and enhanced 18F-FDG uptake in fibroblasts and macrophages. Our results suggested that 18F-FDG accumulation in RA reflects the proliferation of the pannus and inflammatory activity. 18F-FDG PET should be effective for quantifying arthritis activity.

Acknowledgments

We thank Dr. Kazuhiko Yamamoto from the Department of Allergy and Rheumatology, Graduate School of Medicine, The University of Tokyo, for valuable discussions. This work was supported in part by a Health Sciences Research Grant for Research on Specific Diseases from the Ministry of Health, Labour, and Welfare, Japan.

Footnotes

  • Shigeo Koyasu, Department of Microbiology and Immunology, Keio University School of Medicine, 35 Shinanomachi, Shinjuku-ku, Tokyo 160-8585, Japan. koyasu{at}sc.itc.keio.ac.jp

  • COPYRIGHT © 2009 by the Society of Nuclear Medicine, Inc.

References

  1. 1.↵
    Harris ED Jr. Rheumatoid arthritis: pathophysiology and implications for therapy. N Engl J Med. 1990;322:1277–1289.
    OpenUrlCrossRefPubMed
  2. 2.↵
    Lee DM, Weinblatt ME. Rheumatoid arthritis. Lancet. 2001;358:903–911.
    OpenUrlCrossRefPubMed
  3. 3.↵
    Scutellari PN, Orzincolo C. Rheumatoid arthritis: sequences. Eur J Radiol. 1998;27(suppl 1):S31–S38.
    OpenUrlCrossRefPubMed
  4. 4.↵
    Brook A, Corbett M. Radiographic changes in early rheumatoid disease. Ann Rheum Dis. 1977;36:71–73.
    OpenUrlAbstract/FREE Full Text
  5. 5.↵
    Egsmose C, Lund B, Borg G, et al. Patients with rheumatoid arthritis benefit from early 2nd line therapy: 5 year followup of a prospective double blind placebo controlled study. J Rheumatol. 1995;22:2208–2213.
    OpenUrlPubMed
  6. 6.↵
    McQueen FM. Magnetic resonance imaging in early inflammatory arthritis: what is its role? Rheumatology (Oxford). 2000;39:700–706.
    OpenUrlAbstract/FREE Full Text
  7. 7.↵
    Beckers C, Ribbens C, Andre B, et al. Assessment of disease activity in rheumatoid arthritis with 18F-FDG PET. J Nucl Med. 2004;45:956–964.
    OpenUrlAbstract/FREE Full Text
  8. 8.↵
    Polisson RP, Schoenberg OI, Fischman A, et al. Use of magnetic resonance imaging and positron emission tomography in the assessment of synovial volume and glucose metabolism in patients with rheumatoid arthritis. Arthritis Rheum. 1995;38:819–825.
    OpenUrlPubMed
  9. 9.↵
    Beckers C, Jeukens X, Ribbens C, et al. 18F-FDG PET imaging of rheumatoid knee synovitis correlates with dynamic magnetic resonance and sonographic assessments as well as with the serum level of metalloproteinase-3. Eur J Nucl Med Mol Imaging. 2006;33:275–280.
    OpenUrlCrossRefPubMed
  10. 10.↵
    Renner ED, Plagemann PG, Bernlohr RW. Permeation of glucose by simple and facilitated diffusion by Novikoff rat hepatoma cells in suspension culture and its relationship to glucose metabolism. J Biol Chem. 1972;247:5765–5776.
    OpenUrlAbstract/FREE Full Text
  11. 11.↵
    Palmer WE, Rosenthal DI, Schoenberg OI, et al. Quantification of inflammation in the wrist with gadolinium-enhanced MR imaging and PET with 2-[F-18]-fluoro-2-deoxy-d-glucose. Radiology. 1995;196:647–655.
    OpenUrlPubMed
  12. 12.↵
    Goerres GW, Forster A, Uebelhart D, et al. F-18 FDG whole-body PET for the assessment of disease activity in patients with rheumatoid arthritis. Clin Nucl Med. 2006;31:386–390.
    OpenUrlCrossRefPubMed
  13. 13.↵
    Kawamoto T. Use of a new adhesive film for the preparation of multi-purpose fresh-frozen sections from hard tissues, whole-animals, insects and plants. Arch Histol Cytol. 2003;66:123–143.
    OpenUrlCrossRefPubMed
  14. 14.↵
    Feldmann M, Maini RN. Anti-TNF alpha therapy of rheumatoid arthritis: what have we learned? Annu Rev Immunol. 2001;19:163–196.
    OpenUrlCrossRefPubMed
  15. 15.
    Al-Shukaili AK, Al-Jabri AA. Rheumatoid arthritis, cytokines and hypoxia: what is the link? Saudi Med J. 2006;27:1642–1649.
    OpenUrlPubMed
  16. 16.↵
    Gamelli RL, Liu H, He LK, Hofmann CA. Augmentations of glucose uptake and glucose transporter-1 in macrophages following thermal injury and sepsis in mice. J Leukoc Biol. 1996;59:639–647.
    OpenUrlAbstract
  17. 17.↵
    Cornelius P, Marlowe M, Lee MD, Pekala PH. The growth factor-like effects of tumor necrosis factor-alpha: stimulation of glucose transport activity and induction of glucose transporter and immediate early gene expression in 3T3-L1 preadipocytes. J Biol Chem. 1990;265:20506–20516.
    OpenUrlAbstract/FREE Full Text
  18. 18.↵
    Bird TA, Davies A, Baldwin SA, Saklatvala J. Interleukin 1 stimulates hexose transport in fibroblasts by increasing the expression of glucose transporters. J Biol Chem. 1990;265:13578–13583.
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    Taylor PC, Sivakumar B. Hypoxia and angiogenesis in rheumatoid arthritis. Curr Opin Rheumatol. 2005;17:293–298.
    OpenUrlCrossRefPubMed
  20. 20.↵
    Stevens CR, Williams RB, Farrell AJ, Blake DR. Hypoxia and inflammatory synovitis: observations and speculation. Ann Rheum Dis. 1991;50:124–132.
    OpenUrlFREE Full Text
  21. 21.↵
    Peters CL, Morris CJ, Mapp PI, et al. The transcription factors hypoxia-inducible factor 1alpha and Ets-1 colocalize in the hypoxic synovium of inflamed joints in adjuvant-induced arthritis. Arthritis Rheum. 2004;50:291–296.
    OpenUrlCrossRefPubMed
  22. 22.↵
    Kubota R, Yamada S, Kubota K, Ishiwata K, Tamahashi N, Ido T. Intratumoral distribution of fluorine-18-fluorodeoxyglucose in vivo: high accumulation in macrophages and granulation tissues studied by microautoradiography. J Nucl Med. 1992;33:1972–1980.
    OpenUrlAbstract/FREE Full Text
  23. 23.↵
    Yamada S, Kubota K, Kubota R, Ido T, Tamahashi N. High accumulation of fluorine-18-fluorodeoxyglucose in turpentine-induced inflammatory tissue. J Nucl Med. 1995;36:1301–1306.
    OpenUrlAbstract/FREE Full Text
  24. 24.↵
    Lewis JS, Lee JA, Underwood JC, Harris AL, Lewis CE. Macrophage responses to hypoxia: relevance to disease mechanisms. J Leukoc Biol. 1999;66:889–900.
    OpenUrlAbstract
  25. 25.↵
    Simon LM, Robin ED, Phillips JR, Acevedo J, Axline SG, Theodore J. Enzymatic basis for bioenergetic differences of alveolar versus peritoneal macrophages and enzyme regulation by molecular O2. J Clin Invest. 1977;59:443–448.
    OpenUrlPubMed
  26. 26.↵
    Murdoch C, Muthana M, Lewis CE. Hypoxia regulates macrophage functions in inflammation. J Immunol. 2005;175:6257–6263.
    OpenUrlAbstract/FREE Full Text
  27. 27.↵
    Khandwala AS, Gee JB. Factors in glucose oxidation by alveolar macrophages: glucose transport and glycogenolysis. Chest. 1975;67(suppl):60S–63S.
    OpenUrl
  28. 28.↵
    Borregaard N, Herlin T. Energy metabolism of human neutrophils during phagocytosis. J Clin Invest. 1982;70:550–557.
    OpenUrlCrossRefPubMed
  29. 29.↵
    Malone DG, Wahl SM, Tsokos M. Immune function in severe, active rheumatoid arthritis: a relationship between peripheral blood mononuclear cell proliferation to soluble antigens and synovial tissue immunohistologic characteristics. J Clin Invest. 1984;74:1173–1185.
    OpenUrlPubMed
  30. 30.↵
    Shozushima M, Tsutsumi R, Terasaki K, Sato S, Nakamura R, Sakamaki K. Augmentation effects of lymphocyte activation by antigen-presenting macrophages on FDG uptake. Ann Nucl Med. 2003;17:555–560.
    OpenUrlCrossRefPubMed
  • Received for publication November 27, 2008.
  • Accepted for publication February 11, 2009.
Back to top

In this issue

Print
Download PDF
Article Alerts
Email Article
Citation Tools
Bookmark this article

Jump to section