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In permissive tissues, such as the gut and synovium, chronic inflammation can result in the ectopic development of anatomic structures that resemble lymph nodes. These inflammation-induced structures, termed lymphoid neogenesis or tertiary lymphoid organs, may reflect differential stromal responsiveness to the process of lymphoid neogenesis. To investigate the structural reorganization of the microcirculation involved in colonic lymphoid neogenesis, we studied a murine model of dextran sodium sulfate (DSS)-induced colitis. Standard 2-dimensional histology demonstrated both submucosal and intramucosal lymphoid structures in DSS-induced colitis. A spatial frequency analysis of serial histologic sections suggested that most intramucosal lymphoid aggregates developed de novo. Intravital microscopy of intravascular tracers confirmed that the developing intramucosal aggregates were supplied by capillaries arising from the quasi-polygonal mucosal plexus. Confocal optical sections and whole mount morphometry demonstrated capillary networks (185±46um diameter) involving 6–10 capillaries with a luminal diameter of 6.8±1.1um. Microdissection and angiogenesis PCR array analysis demonstrated enhanced expression of multiple angiogenic genes including CCL2, CXCL2, CXCL5, Il-1b, MMP9 and TNF within the mucosal plexus. Intravital microscopy of tracer particle flow velocities demonstrated a marked decrease in flow velocity from 808±901um/sec within the feeding mucosal plexus to 491±155um/sec within the capillary structures. We conclude that the development of ectopic lymphoid tissue requires significant structural remodeling of the stromal microcirculation. A feature of permissive tissues may be the capacity for lymphoid angiogenesis.
In normal circumstances, the peripheral immune system is organized into secondary lymphoid organs such as regional lymph nodes and Peyer's patches (Picker and Butcher, 1992). In some chronic inflammatory diseases, anatomic structures that resemble lymph nodes with B cell follicles and T cell zones form de novo. These inflammation-induced ectopic lymphoid structures have been termed lymphoid neogenesis or tertiary lymphoid organs (Ruddle, 1999; Hjelmstrom, 2001). Some tissues, including the gut, have been associated with lymphoid neogenesis in inflammatory disease states, whereas other tissues, such as the skin, are rarely associated with ectopic lymphoid aggregates (Aloisi and Pujol-Borrell, 2006). These tissue-specific differences suggest the importance of stromal responsiveness to lymphoid neogenesis.
Attempts to investigate stromal adaptation to chronic inflammation have largely focused on endothelial cells. Endothelial cell acquisition of adhesive and chemoattractant properties has been proposed as a mechanism for regulating lymphoid traffic to inflamed tissues (von Andrian and Mempel, 2003). High endothelial venules (HEV), endothelial cells characterized by a cuboidal morphology and a distinctive molecular phenotype (e.g. PNAd+ and CCL21+), are a variable finding in chronically inflamed tissues (Armengol et al., 2001; Weninger et al., 2003; Barone et al., 2005; Manzo et al., 2005). Much of the variability in endothelial adhesive function and chemokine expression may reflect the underlying microvascular architecture. Stromal reorganization, including the adaptive structural change in the microvasculature, may provide an explanation for variability in both endothelial phenotype and tissue responsiveness.
To investigate the structural reorganization of the microcirculation involved in lymphoid neogenesis, we studied a murine model of dextran sodium sulfate (DSS)-induced colitis. Both submucosal and intramucosal lymphoid aggregates were identified in the mouse colon. A frequency analysis suggested that most intramucosal lymphoid aggregates developed de novo. The developing intramucosal lymphoid aggregates were supplied by capillaries arising from the quasi-polygonal mucosal plexus. The time course of aggregate development and the gene expression within the mucosa suggest that the structural changes were the result of inflammation-induced sprouting angiogenesis.
C57B/6 mice (Jackson Laboratory, Bar Harbor, ME), 25–33g, were used in all experiments. The care of the animals was consistent with guidelines of the American Association for Accreditation of Laboratory Animal Care (Bethesda, MD).
In C57/B6 mice, the dextran sodium sulfate (DSS) (TdB Consultancy AB, Uppsala, Sweden) model of colitis was similar to that described previously (Okayasu et al., 1990). Briefly, DSS was freshly prepared and added daily to the drinking water at a final concentration of 5%. The mice were assessed daily for clinical signs and total body weight. The DSS treatment was continued for 5 days then changed to water for the remainder of the experimental period.
Using RFID tagging of each mouse (AVID, www.avidmicrochip.com), body weights and clinical scores were recorded daily. A modification of previously described method (Waidmann et al., 2002), the colitis score incorporated posture (0 normal; 1 abnormal), activity level (0 normal; 1 abnormal), ruffled fur (0 absent; 1 present), rectal prolapse (0 absent; 1 present), feces ((0 normal, 2 liquid, 4 bloody), weight loss (0=<10%, 2=10–20%, 4=>20%). A score of less than 3 was considered minimal or no colitis, 3–5 was moderate colitis, and a score greater than 5 was severe colitis.
As previously described (Ravnic et al., 2007a). The microvideo endoscopy was performed using a multi-purpose rigid endoscope (KSVEA Rigid; 64018 BSA)(Karl Storz, Germany) with a 2.7mm diameter and 18cm length. The rigid optical system included a 30 degree wide angle forward oblique telescope. The KSVEA rigid endoscope used a 175 Watt Xenon light source. The analog video images were digitized for archiving and analysis.
Small mucosal tissue blocks (approximately 10×15mm) were freshly excised from DSS-treated mice, embedded in OCT compound (Miles Labs, Elkhart, IN), and prepared for cryosectioning. Vertical cryosections were prepared in 7–10um thickness slides, stained with hematoxylin and eosin (H&E), and evaluated for lymphoid aggregates. Preliminary microscopic evaluation of each block was performed to ensure an acceptable tissue preparation. In adequately prepared specimens, the number of lymphoid aggregates was assessed based on the H&E staining (Carlsen et al., 2002). Multiple sections were obtained from two parallel regions a minimum of 750um apart; more sections were obtained in regions of apparent confluence so that discrete aggregates could be judged. The histology sections were evaluated with a 500um×750um grid projection by at least two independent observers. The mean aggregate number of these two regions was recorded for each time point.
Immunohistochemistry was performed with commercially available primary antibodies used at a 1:50 concentration. The anti-CD4 (GK1.5, Abcam,, Cambridge UK), anti-CD19 (1D3, BD Pharmigen), anti-CD11b (M1/70, BD Pharmigen, San Jose, CA), F4/80 (CI:A3-1, BD Pharmigen) antibodies were used with a goat anti-rat biotinylated second antibody and developed with neutralite avidin-texas red conjugate (Southern Biotechnology, Birmingham, AL). The biotinylated anti-CD31 (MEC7.46, Cell Sciences, Canton, MA) was developed with the neutralite avidin-texas red conjugate (Southern Biotechnology). The Flk-1/KDR/VEGFR2 antibody (ThermoFisher Scientific) and the anti-CD20 antibody (EP459Y, Abcam) were detected with Qdot 525 goat (F(ab')2 anti-rabbit IgG conjugate (Invitrogen, Eugene, OR).
Cryostat sections were obtained from colon specimens were treated with O.C.T. compound and snap frozen. After warming the slide to 27°C, the sections were fixed for 10 minutes (2% paraformaldehyde and PBS at pH 7.43). The slides were washed with buffer (PBS, 5% sheep serum, 0.1% azide, 1mM MgCl2, 1mM CaCl2) and blocked with 20% sheep serum, 20% goat serum, 0.1% azide in PBS. The slides were treated with monoclonal antibodies (Mab) at 10–20ug/ml. The slides were incubated for one hour at 27°C and washed twice. The detection antibody was added and incubated for 20 minutes at 27°C. The slides were washed twice and examined by fluorescence microscopy.
The exteriorized tissue was imaged using a Nikon Eclipse TE2000 inverted epifluorescence microscope using Nikon objectives of 10×, 20× and 40× linear magnification with infinity correction. An X-Cite (Exfo; Vanier, Canada) 120 watt metal halide light source and a liquid light guide was used to illuminate the tissue samples. Excitation and emission filters (Chroma, Rockingham, VT) in separate LEP motorized filter wheels were controlled by a MAC5000 controller (Ludl, Hawthorne, NY) and MetaMorph software 7.5 (MDS Analytical Technologies, Downingtown, PA). The CFSE tracer (ex 480nm, em 520nm) was imaged with 25nm band pass filters (Omega, Brattleboro, VT). The intravital videomicroscopy 14-bit fluorescent images were digitally recorded on a C9100-02 camera (Hamamatsu, Japan). The C9100-02 camera has a hermetic vacuum-sealed air-cooled head and on-chip electron gain multiplication (2000×). Images with 1000 × 1000 pixel resolution were routinely obtained at 50 fps; frame rates exceeding 50 fps were obtained with binning and subarrays. The images were recorded in image stacks comprising 30 second to 10 minute video sequences.
The structure of the colon microcirculation was characterized by fluorescent vessel painting (Ravnic et al., 2005). After systemic heparinization, the aorta was cannulated and perfused with 15ml of 37°C phosphate buffered saline (PBS) followed by perfusion with a buffered 2.5% glutaraldehyde solution (Sigma). The systemic circulation was perfused with 1,1-dioctadecyl-3,3,3,3-tetramethylindocarbocyanine perchlorate (DiI) or 3,3'-dioctadecyloxacarbocyanine perchlorate (DiO) (10–25ml) as described previously (Ravnic et al., 2005). Immediately following tracer infusion, the organs were harvested, prepared in a 25oC PBS bath, and fixed overnight between glass slides in 4% formalin. After a brief rinse with distilled water, the specimens were stained with DAPI (Vector, Burlingame, CA) and permanently mounted with Vectashield mounting medium (Vector). The fluorescently labeled microvessels were imaged using a Nikon Eclipse TE2000 inverted epifluorescence microscope. Structured illumination confocal microscopy was performed with an Optigrid system (Qioptiq, Rochester, NY)(Lee et al., 2008). The Optigrid uses a one-dimensional optical grid in the form of a Ronchi grating mounted on a piezo-electrically driven actuator. The pattern is moved perpendicular to the grid lines three times producing three separate images that are digitally recombined using a proprietary software algorithm (Volocity 4.4; Improvision, Natick MA).
After euthanasia, subtotal colectomy (ascending to transverse) was performed. The lumen was flushed and opened along the mesenteric border (McDonald and Newberry, 2007). The mucosa was copiously irrigated with cold PBS (4°C) until all debris was removed as determined by stereomicroscopy. The colon wall was immobilized on a standard microscope slide and the mucosa, superficial to the lamina propria, was removed using gentle dissection with a second microscope slide. Limited dissection of the intact superficial (approximately 50um thick) mucosa was confirmed by light microscopy.
Standard RNA isolation procedures were used, including separate laboratory space for tissue harvesting, RNA isolation and PCR processing. Pipets and consumables were regulalrly treated by UV irradiation; work surfaces were routinely cleaned with DNA Exitus Plus (Applichem, Cheshire, CT), standard disinfectants and UV treatment. Routine wipe tests of work areas were performed to screen for nucleic acid contamination.
Total RNA was isolated using Qiagen RNeasy Midi Kit (Qiagen, Valencia, CA). Briefly, the fresh tissue was triturated using a 20g needle until uniformly homogeneous. The tissue lysate was centrifuged at 3000×g for 10minutes and the supernatant (lysate) was removed by pipetting. An equal volume of 70% ethanol was added to lysate and gently mixed. The sample was placed in an RNeasy midi column, centrifuged for 5m at 3000×g and the flow-trough was discarded. After additional RPE buffer was added to the column, the tube was again centrifuged for 5m at 3000×g to dry the RNeasy silica-gel membrane. The RNeasy column was transferred to a collection tube and elution was performed using RNase-free water and centrifugation for 3m at 3000×g. Generally, a second elution step was not performed. Genomic DNA contamination was eliminated by RNase-Free DNase Set (Qiagen). Briefly, 1–2ug of potentially contaminated RNA was treated with DNase buffer, RNase inhibitor and DNase I. In all RNA isolations, the total RNA quality was assessed by using an Agilent 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA). RNA integrity numbers (RIN) (Schroeder et al., 2006) of the RNA samples were uniformally greater than 7.3 (mean 8.5; range 7.3 to 9.8).
First Strand cDNA Synthesis used RT2 First Strand Kit from SuperArray Bioscience Corporation. Mouse Angiogenesis RT2 Profiler PCR Array and RT2 Real-Timer SyBR Green/ROX PCR Mix were purchased from SuperArray Bioscience Corporation (Frederick, MD).
The angiogenesis genes examined in our study include the following (abbreviation, gene name): Angiopoietin 1 (Angpt1, 1110046O21Rik/Ang-1), Angiopoietin 2 (Angpt2, Agpt2/Ang- 2), Chemokine (C-C motif) ligand 2 (CCL2, AI323594/HC11), Collagen, type IV, alpha 3 (Col4a3, [a]3(IV)/alpha3(IV)), Colony Stimulating Factor 3 (granulocyte) (CSF3, Csfg/G-CSF), Chemokine (C-X-C motif) ligand 1 (CXCL1, Fsp/Gro1)), Chemokine (C-X-C motif) ligand 2 (CXCL2, CINC-2a/GROb), Chemokine (C-X-C motif) ligand 5 (CXCL5, AMCF-II/ENA-78), Fibroblast growth factor 1 (Fgf1, Dffrx/Fam), Fibroblast growth factor 2 (Fgf2, Fgf-2/Fgfb), Fibroblast growth factor 6 (Fgf6, Fgf-6), Fibroblast growth factor receptor 3 (Fgfr3, Fgfr-3/HBGFR), Heart and neural crest derivatives transcript 2 (Hand2, AI225906/AI661148), Interferon gamma (Ifng, IFN-g/IFN-gamma), Interleukin 1 beta (Il1b, IL-1beta/Il-1b), Interleukin 6 (Il6, Il-6), Leptin (Lep, ob/obese), Matrix metallopeptidase 9 (MMP9, AW743869/B), T-box 4 (Tbx4, 3930401C23), Transforming growth factor alpha (Tgfa, wa-1/wa1), Transforming growth factor, beta 1 (Tgfb1, TGF-beta1/TGFbeta1), Transforming growth factor, beta 2 (Tgfb2, BB105277/Tgf-beta2), Transforming growth factor, beta 3 (Tgfb3, Tgfb-3), Tumor necrosis factor (TNF, DIF/TNF-alpha), Thymidine phosphorylase (Tymp, 2900072D10Rik/Ecgf1), Vascular endothelial growth factor A(Vegfa, VEGF- A/VEGF120), Vascular endothelial growth factor B (Vegfb, VEGF-B/Vrf), Vascular endothelial growth factor C (Vegfc, AW228853/VEGF-C).
Real-time PCR was performed with SYBR green qPCR master mixes that include a chemically-modified hot start Taq DNA polymerase (SABioscience). PCR was performed on ABI 7300 Real-Time PCR System (Applied Biosystems). For all reactions, the thermal cycling conditions were an initial 50°C for 2 minutes and 95°C for 10 minutes followed by 40 cycles of denaturation at 95°C for 15 seconds and simultaneous annealing and extension at 60°C for 1 minute.
The nanoparticles were developed by Molecular Probes (Invitrogen, Eugene, OR) for intravascular particle tracking (Ravnic et al., 2007b). Characteristics of the particles included superior fluorescence intensity, small size (500nm) and low surface charge content (6.2uEq/gm). The nanoparticles used in this study were green (ex 488; em 510) and infra-red (ex 655nm; em 710nm).
A 5-(and-6)-carboxyfluorescein diacetate, succinimidyl ester (CFSE) (Invitrogen, Eugene, OR) labeling solution was prepared in dimethyl sulfoxide (DMSO) as described (Becker et al., 2004; Ravnic et al., 2006). The freshly prepared CFSE (400ul) was injected into the tail vein of an anesthetized mouse. In some mice, a 10% 250,000kD fluorescein isothiocyanate (FITC)-dextran (Sigma) solution in normal saline was prepared. Mice were injected via the tail vein with 150ul of the prepared solution. The CFSE and FITC-dextran tracers (ex 480nm, em 520nm) were imaged with 25nm band pass filters (Omega).
Tracking of the green and infra-red particles was performed on digitally recorded and distance calibrated multi-image “stacks”(Ravnic et al., 2006). The image stacks produced a sequential time history of velocity and direction as the acquired images were time stamped based on the 100mhz system bus clock of the Xeon processor (Intel, Santa Clara, CA). The movement of individual particles was tracked using the MetaMorph (MDS Analytical Technologies) object tracking applications. The intensity centroids of the particles were identified and their displacements tracked through planes in the source image stack. For displacement reference, the algorithm used the location of the particle at its first position in the stack. Each particle was imaged as a high contrast fluorescent disk and its position was determined with sub pixel accuracy. The image of the particle was tracked using a cross correlation centroid-finding algorithm to determine the best match of the particle/cell position in successive images. With routine distance calibration, the overlay of the image stack provided a quantitative assessment of the particle/cell path. From the XY coordinates, velocity, mean displacement and mean vector length were calculated.
The stream acquired images were stacked to create a time-series of 500 or 1000 consecutive frames. The stacks were systematically analyzed to ensure the absence of motion artifact. The stack "maximum" operation selected the highest intensity value for each pixel location throughout the time-series. The resultant image produced a time-series reconstruction of particle locations during the time interval of the image stack.
Gene expression was calculated using the comparative cycle threshold (Ct) method (Livak and Schmittgen, 2001). Although the data was monitored for nonideal efficiencies, comparable amplification of the target genes and reference genes was assumed. Every effort to optimize the reaction efficiency was made. Validation assays using serial dilutions of the target and reference genes were not routinely performed. The DSS-induced colitis and control data were plotted as a scattergram and a linear regression was calculated with 95% prediction bands after the data was imported into Origin 8.0 (OriginLab, North Hampton, MA). Linear regression was uniformly p<.0001. In nanoparticle velocity analyses, the unpaired Student’s t test for samples of unequal variances was used to calculate statistical significance. The data was expressed as mean ± one standard deviation. The significance level for the sample distribution was defined as P<.01.
Consistent with previous reports (Neurath et al., 2000), the mice in this study (N=92 mice) initially developed weight loss and clinical signs of colitis: the mean weight dropped to 76±6% of baseline on day 7 and gradually recovered to 100±4% of baseline weight on day 28. Similarly, the clinical colitis scores, including ruffled fur, inactivity and diarrhea, peaked on day 8 (score 10±4) and returned to baseline on day 19 (score 0.3±2). Despite the clinical improvement over the first 2–3 weeks, microendoscopy demonstrated ongoing colonic inflammation (Figure 1). Serial histologic sections demonstrated submucosal aggregates that frequently involved both the submucosa and mucosal crypts; these aggregates appeared to span the lamina propria (Figure 1C,D). Relatively smaller intramucosal aggregates were identified within the mucosa; that is, superficial to the lamina propria (Figure 1E,F).
Spatial frequency analysis of the mononuclear aggregates within the submucosa demonstrated an increase in size and prevalence during the 60 day study period (Figure 2, close circles; R=0.79; F=61.8; p<.0001). Similarly, the smaller intramucosal aggregates also increased in size and prevalence (Figure 2, open circles; R=0.62; F=16.4; p<.0001). As expected, immunophenotyping demonstrated T-cell (CD4) and B-cell (CD19) expression within the submucosal aggregates (Figure 3A–C). The prevalence of B cells within the aggregates, as demonstrated by anti-CD19 and anti-CD20 staining, increased in frequency between 30 and 60 days. The increase in B-cell frequency, demonstrated by both immunofluorescence and H&E staining, was associated with an increasing frequency of lymphoid follicles. Although most intramucosal and submucosal aggregates were phenotypically similar, 6–10% of the intramucosal mononuclear aggregates demonstrated a predominance of monocytoid markers (CD11b and F4/80)(Figure 3D–F).
The development of intramucosal lymphoid tissue suggested the possibility of structural changes in the mucosal microcirculation. To investigate changes in the vascular microarchitecture, the mucosal plexus was examined using intravital videomicroscopy and fluorescent vessel painting. In control mice, the mucosal plexus--a quasi-polygonal network of vessels surrounding the mucosal crypts--was a continuous plexus without a specialized vascular supply to lymphoid tissue. In contrast, 21 to 60 days after the onset of inflammation, intravital microscopy studies of the mucosal plexus demonstrated distinctive microcirculatory structures composed of a small network of capillaries. The structures were 185±46um (N=12) in diameter and appeared to be contiguous with the mucosal plexus (Figure 4). Morphometry based on fluorescent vessel painting and 3D tissue mounts of the capillaries indicated a microvessel diameter of 6.8±1.1um (N=6). Consistent with the histologic analysis, the rarity of similar structures in control mice suggested that the capillary structures developed de novo. To explore the gene expression potentially involved in sprouting angiogenesis, mRNA was isolated from microdissected mucosal plexus in DSS-induced colitis and control mice. The expression of genes implicated in angiogenesis was explored using the angiogenesis pathway PCR arrays at 4 timepoints after the induction of DSS colitis: 7, 14, 31 and 65 days (Figure 5). The expression of CXCL2, Il-1b, CXCL5, CCL2, TNF and MMP9 peaked at 14 days after the onset of DSS-induced colitis (Figure 5B). In this bulk RNA, angiogenic factors with a less notable inflammatory association, such as Angpt1, Angpt2, Vegfa, Vegfb, and Vegfc, were not significantly elevated relative to controls (Figure 6).
The functional implications of the lymphoid angiogenesis was investigated by fluorescent intravital videomicroscopy. In the chronic phase, 30 to 60 days after the onset of chemically-induced colitis, intravenously injected fluorescent nanoparticles were tracked through the mucosal capillary structures (Figure 7A,B). Nanoparticle flow demonstrated that the particles passed directly from the mucosal plexus into the capillary structures, confirming both structural and functional continuity. Frequently, the particles exited the capillary structures and passed into deeper collecting veins. The flow through the mucosal plexus structures was notable for a significant decrease in flow velocity when compared to the feeding vessels within the mucosal plexus (Figure 7C–E). The analysis of intravital microscopy recordings (N=6 mice) showed that particles passing into these de novo vessels demonstrated a mean velocity of 491±155um/sec. In contrast, the feeding vessels of the mucosal plexus demonstrated a mean velocity of 808±901um/sec (Figure 7F: p<.01).
In this report, we studied the microvascular adaptations associated with prolonged inflammation in DSS-induced murine colitis. Although both submucosal and intramucosal lymphoid aggregates were identified, the development of lymphoid neogenesis within the superficial mucosa was associated with structural reorganization of the microcirculation. A frequency analysis suggested that most intramucosal lymphoid aggregates developed de novo. The intramucosal aggregates were supplied by capillaries arising from the quasi-polygonal mucosal plexus. The expression of genes associated with both inflammation and angiogenesis suggested that the structural changes were the result of inflammation-induced sprouting angiogenesis.
The structural changes observed in this study highlight the importance of the mucosal stroma in sustaining a peripheral immune response. Previous work has focused on the plasticity of vascular endothelial cells in adapting to peripheral inflammation. Endothelial cells can undergo dramatic inflammation-induced changes in morphology--from flat conduit lining cells to cuboidal high endothelial venule (HEV) cells (Freemont, 1988; Sasaki et al., 1994; Peng, 1996). On a molecular level, the stromal-endothelial interactions in ulcerative colitis and rheumatoid arthritis can stimulate the expression of PNAd, CCL21, and CXCL13 proteins (Takemura et al., 2001; Salomonsson et al., 2003; Carlsen et al., 2004; Manzo et al., 2005). Consistent with other studies of inflammation (Weninger et al., 2003), PNAd expression in our model was inconsistent (not shown) suggesting that the variability in HEV morphology and PNAd expression may reflect different stages of ectopic lymphoid aggregate development. Regardless, the significant structural remodeling--including the apparent sprouting growth of a complex arrangement of capillaries--suggests that stromal adaptations play an important role in the pathophysiology of prolonged DSS-induced colitis.
Our exploratory analysis of mRNA expression within the mucosal plexus demonstrated several mediators previously associated with angiogenesis. Three members of the CXC chemokine family, known to promote angiogenesis (Strieter et al., 2005), were expressed at high levels during the peak of the inflammation. mRNA from the CXCL1, CXCL2 and CXCL5 genes were expressed at levels 90-to 8000-fold greater than controls. These CXC family chemokines signal through the CXCR2 receptor—a receptor that has been implicated in vivo in models of corneal neovascularization (Addison et al., 2000) and wound repair (Devalaraja et al., 2000). The association of MMP9 with tissue remodeling (Page-McCaw et al., 2007) suggests a functional role for extracellular proteases in the remodeling necessary for the development of both lymphoid aggregates and lymphoid angiogenesis. Furthermore, the inflammatory mediators IL-1b and TNF also have been implicated in angiogenesis (Maruotti et al., 2006). In contrast, the expression of angiogenic factors such as Angpt1 and Vegfa were not elevated relative to controls. This finding may reflect the bulk sampling of mRNA. Although the mucosal plexus was microdissected from the remainder of the colon wall, the samples included bulk mRNA from the the perivascular inflammatory cells as well as the mucosal plexus vessels. Discrete spatial sampling, enabled by laser capture microdissection, may be necessary to elucidate the participation of these factors.
Gut-associated lymphoid tissue has been separated into effector sites which consist of lymphocytes scattered throughout the superficial mucosal tissue and the induction sites present in organized lymphoid tissues (Mowat, 2003; Spahn and Kucharzik, 2004). The inductive sites include Peyer's patches, mesenteric lymph nodes, and isolated lymphoid follicles. The contemporary understanding of immunologic function is that antigen presentation and the generation of antigen-specific effector cells occurs in inductive tissues, and that effector cells migrate into superficial mucosal tissues. Our observation of progressive organization of the superficial mucosal compartment suggests that this initial functional distinction may evolve during the subacute phase of inflammation leading to the presence of both inductive and effector elements with the chronically inflamed colonic mucosa. This functional evolution within mucosal tissue is suggested by human studies of secondary and ectopic lymphoid tissue (Manzo et al., 2007). Distorted crypt architecture, intramucosal inflammatory cells, and lymphoid aggregates were features present in 79% of ongoing inflammatory bowel disease and were highly predictive of chronic colitis (Surawicz and Belic, 1984)
The development of inductive sites may also provide an explanation for the spatial distribution of the intramucosal aggregates. The sporadic distribution of intramucosal lymphoid aggregates does not reflect any vascular microarchitectural feature that might predispose to capillary sprouting and lymphoid angiogenesis. Rather than a structural predisposition to lymphoid-associated angiogenesis, we suspect that the spatial distribution of lymphoid aggregates may reflect the location of antigen presenting cells, such as dendritic cells, in the initiation of lymphoid neogenesis (Carragher et al., 2008). The importance of antigen presenting cells in lymphoid neogenesis may also help explain the occasional concentration of monocytoid cells within the intramucosal aggregates.
An advantage of our study was the use of intravascular tracers to demonstrate structural continuity between the capillary structures and the mucosal plexus. Because the particles were inert and charge-neutral, they could be tracked through the microcirculation without the concern of unanticipated biomolecular interactions with vascular lining cells. The particles provided a useful measure of both flow velocity and network flow fields within the sprouting capillary structures. An interesting observation was the diminished flow velocity within the capillary structures; velocities were sufficiently diminished to be within the physiologic range of rolling velocities in secondary lymphoid tissue (Stein et al., 1999). Thus, even if some of the typical secondary lymphoid organ receptor-ligand interactions were not present, the microhemodynamic conditions were nonetheless suitable for lymphoid adhesion and transmigration (Li et al., 2001).
Finally, an assessment of capillary structure and intravacular tracer velocity permits an assessment of blood flow within the mucosal aggregates. Assuming an average of 10 capillaries with cylindrical geometry, and a cardiac output of 20ml/min (Janssen et al., 2002), the perfusion of an average capillary structure would be approximately 0.0015% of cardiac output; that is, approximately 10% of the perfusion of a popliteal lymph node in the sheep (0.014% cardiac output) or a comparable node in the rabbit (0.011%)(Hay and Hobbs, 1977). This finding indicates that even small and developing lymphoid structures possess the potential for a substantial exposure to circulating lymphocytes and a significant contribution to the peripheral immune response.
Supported in part by NIH Grant HL47078 and HL75426