To develop an in vivo imaging method to assess lymphatic draining function in the K/BxN mouse model of inflammatory arthritis.
To develop an in vivo imaging method to assess lymphatic draining function in the K/BxN mouse model of inflammatory arthritis.
Indocyanine green, a near-infrared fluorescent dye, was injected intradermally into the footpads of wild-type mice, mouse limbs were illuminated with an 806-nm near-infrared laser, and the movement of indocyanine green from the injection site to the draining popliteal lymph node (LN) was recorded with a CCD camera. Indocyanine green near-infrared images were analyzed to obtain 5 measures of lymphatic function across time. Images of K/BxN arthritic mice and control nonarthritic littermates were obtained at 1 month of age, when acute joint inflammation commenced, and again at 3 months of age, when joint inflammation became chronic. Lymphangiogenesis in popliteal LNs was assessed by immunochemistry.
Indocyanine green and its transport within lymphatic vessels were readily visualized, and quantitative measures were derived. During the acute phase of arthritis, the lymphatic vessels were dilated, with increased indocyanine green signal intensity and lymphatic pulses, and popliteal LNs became fluorescent quickly. During the chronic phase, new lymphatic vessels were present near the foot. However, the appearance of indocyanine green in lymphatic vessels was delayed. The size and area of popliteal LN lymphatic sinuses progressively increased in the K/BxN mice.
Our findings indicate that indocyanine green near-infrared lymphatic imaging is a valuable method for assessing the lymphatic draining function in mice with inflammatory arthritis. Indocyanine green–near-infrared imaging of K/BxN mice identified 2 distinct lymphatic phenotypes during the acute and chronic phase of inflammation. This technique can be used to assess new therapies for lymphatic disorders.
Inflammatory erosive arthropathies, such as rheumatoid arthritis (RA), are characterized by pannus formation and joint tissue damage. While it is known that inflammation in RA induces angiogenesis and tissue edema due to the accumulation of fluid leaking from newly formed blood vessels into the interstitial space (1) and that this interstitial fluid contains high levels of cytokines and chemokines that may contribute to joint tissue destruction (2), little is known about the compensatory lymphatic mechanisms responsible for the removal of accumulated fluid from the joint interstitial space that attenuate local edema, inflammation, and bone and cartilage erosion. Since this is largely due to a lack of physiologic outcome measures to assess lymphatics and lymph flow during arthritis, novel approaches to the quantification of this central biomarker of disease pathogenesis are warranted.
The lymphatic system drains interstitial fluid to local draining lymph nodes (LNs) through lymphatic vessels (3). The lymph generated in RA joints and the lymphatic flow are increased during disease progression and contain significantly elevated levels of proinflammatory cytokines and chemokines (4). Enzyme-linked immunosorbent assay of synovial fluid from RA patients demonstrated elevated levels of vascular endothelial growth factor C (VEGF-C), the primary lymphatic growth factor, which correlated significantly with tumor necrosis factor (TNF) and interleukin-1 levels in RA (5). Immunostaining of synovial tissue from patients with RA and osteoarthritis using antibodies that recognize lymphatic endothelial cells or lymphatic growth factors demonstrated increased lymphatic vessel formation (6–8). We recently reported that the blockade of lymphangiogenesis at the beginning of arthritis development increased the severity of joint tissue injury (9). Thus, lymphatic growth factor expression, lymphatic vessel formation, and lymph flow are all increased in RA joints, suggesting that increased lymphangiogenesis is a compensatory mechanism to ameliorate inflammatory arthritis. However, the role of the lymphatic system in the pathogenesis of inflammatory arthritis has not been elucidated because of inadequate imaging modalities for assessing lymphatic function in murine models.
Several imaging methods have been used in rodents to measure lymph flow, including whole-body radiography, magnetic resonance imaging, optical imaging, and intravital microscopy (10–15). Although these technologies can visualize the path of a contrast agent within lymphatic vessels, they are expensive and require special instruments and highly trained personnel, thus limiting routine use in laboratory animals. Indocyanine green is a dye that is safe when administered as an intravenous bolus and is used for multiple purposes (16). Once injected, it stays in the blood and is excreted in bile without being reabsorbed from the gut. Although it is green, indocyanine green has absorption and emission spectra within the near-infrared range, enabling deep-tissue penetration and fluorescence visualization. Sharma et al (17) used indocyanine green near-infrared fluorescence to examine the lymphatics in swine. They also completed a phase I clinical trial using small doses of indocyanine green to assess lymph traffic from tumor to sentinel LNs in breast cancer (18). However, this method has not been used in laboratory mouse models of arthritis.
In the present study, we used indocyanine green near-infrared lymphatic imaging to examine progressive changes in lymphatic function in the legs of K/BxN mice (19), a murine model of inflammatory arthritis. During the acute phase of arthritis, lymphatic flow increased, resulting in dilated lymphatic vessels and severe ankle joint swelling. During the chronic phase, new lymphatic vessels with decreased lymph flow developed in the foot. Indocyanine green near-infrared imaging demonstrated impaired lymphatic function in inflamed joints, suggesting that the improvement in local lymph flow may represent a new target for RA therapy in patients with severe disease. Additionally, we found that indocyanine green near-infrared imaging is an effective method for assessing lymphatic function in vivo.
KRN/NOD-transgenic mice were obtained by crossing KRN-transgenic males (B6 genetic background; provided by Drs. C. Benoist and D. Mathis, Harvard Medical School, Boston, MA) with female nonobese diabetic (NOD) mice (NOD/LtJ; The Jackson Laboratory). (KRN/NOD)F1 offspring mice were bled on day 21 after birth, and those expressing the αVβ6 T cell receptor transgene were identified by flow cytometry. These were the K/BxN mice. F1 offspring mice that did not express the transgene were used as controls. All K/BxN mice develop severe ankle joint inflammation at ∼1 month of age. For flow cytometry, whole blood was incubated on ice with phycoerythrin (PE)–conjugated monoclonal anti-mouse CD4 T cell (0.5 mg) and fluorescein isothiocyanate (FITC)–conjugated monoclonal anti-mouse αVβ6 T cell receptor β-chain (0.5 mg) antibodies (BD PharMingen). Animals that carried CD4+αVβ6+ cells were identified as KRN-transgene positive.
For indocyanine green near-infrared imaging, animals were anesthetized with 2% isoflurane (Butler Animal Health Supply) and maintained with 1% isoflurane in oxygen. Animals underwent indocyanine green near-infrared imaging at 1 and 3 months of age. The University of Rochester Medical Center Institutional Animal Care and Use Committee approved all animal studies. For determination of the severity of edema, the ankle thickness (mm) was measured at 1 month and 3 months of age using calipers.
A Spy1000 system (Novadaq Technologies) was used. This system consists of an 806-nm laser providing maximum illumination intensity that are of ∼30 mW/cm2 at a fixed focal length. This instrument was modified to provide continuous operation and adjustment of laser intensity and was equipped with a second PAL camera. Filters permitted the measurement of light above 815 nm. The video outputs of the camera were attached to the network using an Axis 241SA video server that was monitored using SecuritySpy (Ben Bird; online at www.bensoftware.com) and converted motion JPEG image streams into QuickTime movies (Apple Computers). Individual JPEG image sequences were then exported and read for further analysis with ImageJ (NIH Image, National Institutes of Health; online at http://rsbweb.nih.gov/ij/).
Indocyanine green (Acorn or Pulsion) was dissolved in distilled water at 0.1 μg/μl, and 10 μl (1 μg) of the green solution was injected intradermally into the mouse footpad using a 30-gauge needle. Before indocyanine green injection, fur was removed from the legs with hair removal lotion. Animals were placed in a recumbent position, and their legs were restrained with tape on an isothermal gel pad during imaging sessions. Indocyanine green fluorescence was recorded for 1–2 hours immediately and again for 5 minutes 24 hours after indocyanine green injection. The QuickTime movies were examined, and the time between footpad injection and the appearance of indocyanine green in the popliteal LNs was recorded. Sequential images from the movie file were exported, and the intensity of indocyanine green fluorescence of popliteal LNs and footpads was determined using ImageJ software. A fixed circular region of interest (ROI) was drawn over the vessels or popliteal LNs and nearby background tissue. The signal intensity was defined as the signal intensity minus the background intensity. The following outcome measures were derived from the indocyanine green near-infrared images: 1) T-initial (T-in), which is the time that it takes for indocyanine green to be detected in vessels; 2) S-max, which is the maximum indocyanine green signal intensity observed in the popliteal LN during the first day imaging session; 3) T-max, which is the time that it takes for a popliteal LN to achieve maximal indocyanine green signal intensity; 4) percent clearance, which is an assessment of indocyanine green washout through the lymphatics and is quantified as the percent difference in indocyanine green signal intensity between the 2 indocyanine green near-infrared images from the ROI of the popliteal LNs or footpad at S-max (first day) and 24 hours after indocyanine green injection; and 5) pulse, which is the number of indocyanine green pulses that pass the ROI within 400 seconds.
Popliteal LNs were frozen in liquid nitrogen and embedded in OCT matrix. Frozen sections (10 μm) were stained with a mixture of FITC-conjugated rabbit anti–lymphatic vessel endothelial receptor 1 (LYVE-1) antibody followed by Alexa Fluor 488 goat anti-rabbit IgG and PE-conjugated anti-CD31 antibody, as previously described (8). Four pictures (at 10× magnification) were taken from each section from different fields. Lymphatic vessels were quantified by a point counting method, as previously described (20). For each mouse, 2 sections were cut at 250 μm apart, and the area and size of LYVE-1+ lymphatic vessels were measured for the entire popliteal LN and expressed as the percentage of LYVE-1+ vessels per popliteal LN, as previously described (8). Data are presented as the mean ± SD of 4–6 popliteal LNs.
All results are presented as the mean ± SD. Comparisons between 2 groups were analyzed using Student's unpaired 2-tailed t-test. One-way analysis of variance and Dunnett's post hoc multiple comparisons were used for comparisons among ≥3 groups. P values less than 0.05 were considered significant.
To establish a quantitative indocyanine green near-infrared lymphatic imaging protocol for the mouse leg, we first evaluated the clearance behavior of indocyanine green throughout the animal. Indocyanine green was injected intradermally into the footpad, and the path of indocyanine green clearance through 2 parallel lymphatic vessels in the lower limb to the popliteal LN was clearly visible by near-infrared imaging (Figures 1A and B). Whole-body indocyanine green near-infrared imaging demonstrated that the fluorescent dye migrated from the popliteal LN to the gluteal and iliac LNs (Figure 1C). From there it was efficiently cleared through the digestive system, such that no detectable indocyanine green remained in wild-type mice 48 hours after injection. These findings are consistent with those of previous lymphatic tracer studies (21) and confirm that measurement of indocyanine green flow between the foot and the popliteal LN reflects lymphatic function in the lower limbs of mice.
To quantify lymphatic function, ROI were retrospectively identified on the indocyanine green near-infrared images from the 1–2-hour real-time video to quantify 5 independent metrics (Figure 2A). As described above, these metrics were T-in, S-max, T-max, lymphatic pulse (amplitude of signal intensity and time interval between pulses), and rate of indocyanine green clearance from the injection site in the foot (percent indocyanine green signal at 24 hours). Figure 2B shows a Z score data set from a real-time indocyanine green near-infrared video analyzed over a 10-minute period during which T-in was achieved. The Z score was calculated as follows: (current observation − mean)/SD. The Z scores were plotted against the frame number (time), where a Z score >3 (P < 0.01) was considered to be T-in time. Figure 2C illustrates the mean fluorescence intensity profile of the ROI for lymphatic pulse over time, which shows that the lymphatic vessels in the leg of a wild-type mouse under deep anesthesia beat every 55 seconds. This is in dramatic contrast to the mouse's vascular pulse, which is driven by a heart rate of ∼550–600 beats per minute. These data also draw attention to a common misconception that lymph migration is achieved via adjacent muscle tissue contraction, when in fact the indocyanine green migration in our system was entirely dependent on lymphatic vessel contraction, which contains its own pulse.
As a prelude to our studies examining the effects of altered lymphatic flow on the severity of joint inflammation, we first excluded the possibility that indocyanine green footpad injection itself causes local inflammation. Injection of indocyanine green into the footpad of wild-type mice did not result in histologic changes at the injection sites on hematoxylin and eosin–stained sections (results not shown). We then examined the progress of ankle synovitis in K/BxN mice during the acute and chronic phases of arthritis by comparing paw thickness in the same animals at 1 and 3 months of age and found that foot swelling decreased with age (mean ± SD ankle thickness 4.5 ± 0.5 mm at 1 month versus 3.3 ± 0.2 mm at 3 months; P < 0.05). Taken together, these findings demonstrate that our indocyanine green near-infrared experimental approach is innocuous, and that 1-month-old K/BxN mice display acute ankle synovitis, which is abated during the chronic phase of arthritis as seen in 3-month-old K/BxN mice.
To examine the changes in lymphatic function during the natural history of inflammatory arthritis, we performed indocyanine green near-infrared imaging on the legs of K/BxN arthritic mice and control nonarthritic littermates prospectively. K/BxN mice develop severe inflammatory arthritis at ∼1 month of age. Joint redness and swelling occur within 2–3 days during the acute phase. Afterward, inflammation persists into the chronic phase (>3 months old), during which the redness and swelling decline but the tissue damage remains (22, 23).
The indocyanine green near-infrared images showed that control mice had 1 or 2 major lymphatic vessels linking the foot area to the popliteal LN. These vessels were thin, with weak indocyanine green fluorescence (arrows in Figure 3A). Accordingly, the S-max was low, and the T-max was ∼30–50 minutes. There were no notable differences in lymphatic parameters between 1-month-old and 3-month-old control mice (Figure 3A). These observations suggest a consistent level of lymph movement from the foot area to the popliteal LN in the normal mouse leg under anesthesia. In contrast, K/BxN arthritic mice had a dramatically different lymphatic phenotype. During the acute phase (at 1 month of age), extensively dilated and irregular lymphatic vessels (arrowheads in Figure 3A) were observed, and a very bright indocyanine green signal was present in vessels and popliteal LNs. During the chronic phase, the indocyanine green signal intensity and the size of the lymphatic vessels returned to control levels. However, multiple newly formed lymphatic vessels near the foot area were visualized (arrowheads in Figure 3B).
These observations were confirmed by quantitative analysis (Figure 3C). During the acute phase of arthritis in K/BxN mice, the T-in was significantly decreased (13-fold), and the S-max was significantly increased (10-fold). In contrast, during the chronic inflammation phase in 3-month-old K/BxN mice, the T-in, S-max, and indocyanine green clearance were all similar to the values in their nontransgenic littermates. However, the lymphatics in the legs of K/BxN mice with chronic arthritis were still abnormal, as evidenced by a 2-fold increase in T-max, which was also significantly greater than that seen in 1-month-old K/BxN mice with acute arthritis. Taken together, these data predict that the lymphatic pulse is significantly increased during the acute phase, to achieve accelerated indocyanine green migration. However, during chronic arthritis, this accelerated pulse decreases and lymphangiogenesis predominates to increase the number and volume of lymphatic vessels.
To evaluate the effects of acute and chronic ankle arthritis on the lymphatic pulse in the leg, we performed indocyanine green near-infrared analyses as shown in Figure 2C. We found that during the acute phase of arthritis, K/BxN mice had a significantly increased pulse rate, reaching ∼5 pulses per minute (Figure 4). In contrast, the lymphatic pulses in the chronic phase returned to control levels (∼1 pulse per minute). Thus, these data support a model of increased lymphatic flow in acute inflammation, followed by increased lymphangiogenesis with decreased lymphatic flow during the progression of inflammatory arthritis.
Since our lymphatic flow data clearly predicted significant changes in the number of lymphatic vessels and overall lymphatic volume of the lower limb during arthritis progression in K/BxN mice, we examined the lymphatic vasculature in popliteal LNs by immunostaining with anti–LYVE-1 antibody to identify lymphatic sinuses, as previously described (9, 24). The K/BxN popliteal LNs had significantly increased volume and weight at 1 month, which increased to 10 times that of control mice by 3 months (Figure 5A). Compared with control popliteal LNs, popliteal LNs from 1-month-old K/BxN mice had increased numbers of LYVE-1+ lymphatic sinuses with narrow lumens (Figure 5B, parts a, c, and e). Popliteal LNs from 3 month-old K/BxN mice had even more LYVE-1+ lymphatic sinuses, the majority of them having extensive and dilated lumens (indicated by the arrowheads in Figure 5B, parts d and f). Findings from immunostaining were confirmed by histomorphometric analysis (Figure 5C). We also performed immunohistochemical analysis for CD31+ blood vessels and found that angiogenesis was also increased in K/BxN popliteal LNs as compared with control LNs, but to a lesser degree than lymphangiogenesis (Figure 5B and results not shown).
Although there have been significant advances in our understanding of the etiology and pathogenesis of RA, several enigmatic aspects of this disease remain to be elucidated. One of them is the manner in which the disease waxes and wanes over long periods of time and the sudden onset of exacerbated joint inflammation and pain commonly referred to as arthritic flare. Since repeated efforts to identify changes in autoimmunity have failed to produce significant findings, it appears more likely that these varying disease states are caused by environmental and epigenetic influences on a chronic condition. One mechanism by which these influences could dramatically affect RA is by altering the efferent lymphatic flow from the joint, which functions to perpetually remove immune cells and catabolic factors that would otherwise accumulate, causing swelling, connective tissue destruction, and pain. Unfortunately, the absence of a minimally invasive outcome measure to quantify lymphatic flow has prevented this area of study from moving forward, such that the direct role of lymphatic drainage on RA pathogenesis remains largely unknown. Thus, in order to overcome this obstacle and to better understand the relationship between lymphatic flow and inflammatory arthritis progression, we established an indocyanine green near-infrared lymphatic imaging protocol in mouse legs to visualize and quantify changes during the natural history of inflammatory arthritis in K/BxN mice.
In our longitudinal studies, we observed 2 distinct lymphatic phenotypes during the acute versus chronic phases of arthritis in K/BxN mice. At the onset of severe ankle inflammation, lymphatic flow was dramatically increased, as evidenced by a 5-fold increase in the lymphatic pulse and several quantitative measures of indocyanine green migration in lymphatic vessels. We also observed that the lymphatic vessels appeared to be extremely dilated and very bright, indicating that a large amount of lymph was entering afferent lymphatic vessels within a short time as a consequence of acute inflammation, where fluid is rapidly accumulating in the interstitial space due to blood vascular leakage (25). This is consistent with the clinical observation of increased lymph flow in lymphatic vessels that drain inflamed joints in RA patients (4). However, this increase in lymphatic flow is not sustained, and the joint disease progresses to a chronic phase, which is characterized by numerous new lymphatic vessels that drain the inflamed tissue, with a decreased lymphatic pulse and slower lymphatic flow. The notion of this de novo lymphangiogenesis is supported by studies demonstrating increased lymphatic vasculature in the synovium of RA patients and animals by immunohistochemistry (6–8). In other models of inflammation, such as bacteria-induced lung inflammation, newly formed lymphatic vessels are also observed (26, 27).
These observations raise the question of why there is slower lymphatic flow during the chronic phase, when the large number of newly formed lymphatic vessels should be able to increase lymphatic flow. The simplest explanations are either that the vessels are immature and leaky or that they are nonfunctional. Interestingly, it has been reported that transgenic overexpression or administration of VEGF-A or VEGF-C induces lymphatic vessel formation, but these newly formed vessels leak (28, 29). However, we did not observe indocyanine green fluorescence around the lymphatic vessels, suggesting that it is unlikely that the slower lymph flow is due to vessel leakage. As such, our attention has turned toward understanding the mechanisms responsible for the lymphatic pulse, with the goal of identifying factors that could be pharmacologically manipulated to increase lymphatic flow and decrease joint inflammation.
Our studies suggest that there is a direct relationship between dysregulated lymph flow and inflammation. This theory is now supported by 2 lines of experimental evidence in mouse models of inflammatory arthritis. First, loss-of-function studies have shown that inhibition of lymphangiogenesis with anti–VEGF receptor 3 neutralizing antibodies decreases lymphatic flow from the foot and increases the severity of ankle joint inflammation (9). Second, we have recently completed a series of gain-of-function studies in which lymphangiogenesis was induced via intraarticular injection of recombinant adeno-associated virus that overexpresses VEGF-C, which significantly increased lymph flow and reduced inflammation in arthritic joints (Zhou Q, et al: unpublished observations). These data support the hypothesis of lymphatic involvement in RA. The lymphatic machinery is composed of lymphatic endothelium, muscle (30), and valve (31). Currently, it is not known whether chronic joint inflammation in RA patients is associated with a defect in any of these components. Similarly, very little is known about the cellular and molecular mechanisms controlling the function of lymphatic endothelium, muscle, and valve under normal and arthritic conditions. By using indocyanine green near-infrared imaging of lymphatic flow in animal models and clinical studies, it may be possible to elucidate the mechanisms responsible for arthritic flares and to develop therapeutic interventions to prevent and treat RA progression.
All authors were involved in drafting the article or revising it critically for important intellectual content, and all authors approved the final version to be published. Dr. Xing had full access to all of the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.
Study conception and design. Zhou, Wood, Schwarz, Wang, Xing.
Acquisition of data. Zhou, Wood, Xing.
Analysis and interpretation of data. Zhou, Wood, Schwarz, Wang, Xing.
The authors would like to thank Drs. C. Benoist and D. Mathis (Harvard Medical School, Boston, MA) for providing male KRN-transgenic mice, Dr. B. Boyce for helpful discussion, and Ms Yanyun Li for technical assistance with the histology. Dr. D. Golijanin provided advice, encouragement, and access to the Spy1000 system obtained under a research agreement between the University of Rochester and Novadaq Technologies.