Microvascular blood flow and stasis in transgenic sickle mice Utility of a dorsal skin fold chamber for intravital microscopy.код для вставкиСкачать
American Journal of Hematology 77:117–125 (2004) Microvascular Blood Flow and Stasis in Transgenic Sickle Mice: Utility of a Dorsal Skin Fold Chamber for Intravital Microscopy Venkatasubramaniam S. Kalambur,1 Hemchandra Mahaseth,2 John C. Bischof,1 Miroslaw C. Kielbik,2 Thomas E. Welch,2 Åsa Vilbäck,2 David J. Swanlund,1 Robert P. Hebbel,2,3 John D. Belcher,2,3 and Gregory M. Vercellotti2,3* 1 Department of Mechanical Engineering, University of Minnesota, Minneapolis, Minnesota 2 Department of Medicine, University of Minnesota, Minneapolis, Minnesota 3 Vascular Biology Center, University of Minnesota, Minneapolis, Minnesota Vascular inflammation, secondary to ischemia–reperfusion injury, may play an essential role in vaso-occlusion in sickle cell disease (SCD). To investigate this hypothesis, dorsal skin fold chambers (DSFCs) were implanted on normal and transgenic sickle mice expressing human a and bs/bs-Antilles globin chains. Microvessels in the DSFC were visualized by intravital microscopy at baseline in ambient air and after exposure to hypoxia–reoxygenation. The mean venule diameter decreased 9% (P < 0.01) in sickle mice after hypoxia–reoxygenation but remained constant in normal mice. The mean RBC velocity and wall shear rate decreased 55% (P < 0.001) in sickle but not normal mice after hypoxia–reoxygenation. None of the venules in normal mice became static at any time during hypoxia–reoxygenation; however, after 1 hr of hypoxia and 1 hr of reoxygenation, 11.9% of the venules in sickle mice became static (P < 0.001). After 1 hr of hypoxia and 4 hr of reoxygenation, most of the stasis had resolved; only 3.6% of the subcutaneous venules in sickle mice remained static (P = 0.01). All of the venules were flowing again after 24 hr of reoxygenation. Vascular stasis could not be induced in the subcutaneous venules of sickle mice by tumor necrosis factor alpha (TNF-a). Leukocyte rolling flux and firm adhesion, manifestations of vascular inflammation, were significantly higher at baseline in sickle mice compared to normal (P < 0.01) and increased 3-fold in sickle (P < 0.01), but not in normal mice, after hypoxia–reoxygenation. Plugs of adherent leukocytes were seen at bifurcations at the beginning of static venules. Misshapen RBCs were also seen in subcutaneous venules. Am. J. Hematol. 77:117–125, 2004. ª 2004 Wiley-Liss, Inc. Key words: sickle cell disease; intravital microscopy; skin fold chambers; blood flow; stasis; vaso-occlusion INTRODUCTION Sickle cell disease (SCD) is characterized by recurring acute vaso-occlusive episodes resulting in damage to multiple organs . The substitution of valine for glutamate at the sixth position of the b chain of hemoglobin  results in the polymerization of sickle hemoglobin and the sickling of RBCs under deoxygenated conditions. The polymerization of deoxygenated sickle hemoglobin is the primary event in the molecular pathogenesis of SCD and is responsible for the vaso-occlusive phenomena that are the hallmarks of the disease . The repetitive vaso-occlusive episodes cause tissue ischemia and ª 2004 Wiley-Liss, Inc. Contract Grant Sponsor: NHLBI; Contract grant numbers: HL67367, HL55552 *Correspondence to: Gregory M. Vercellotti, M.D., University of Minnesota, Department of Medicine, Division of Hematology, Oncology and Transplantation, 420 Delaware St. SE, MMC 480, Minneapolis, MN 55455. E-mail: firstname.lastname@example.org Received for publication 12 May 2003; Accepted 13 April 2004 Published online in Wiley InterScience (www.interscience.wiley.com). DOI: 10.1002/ajh.20143 118 Kalambur et al. reperfusion injury, resulting in endothelial cell activation and inflammation [4,5]. Sickle patients have multiple indicators of an inflammatory response including elevated white counts [6–9], C-reactive protein levels [10–13], cytokines [14–17], as well as activated monocytes [13,18], neutrophils [19–21], platelets [18,22–28], and endothelial cells [29,30] in circulation. Transgenic mice expressing human a and bS globins provide unique models to study abnormalities in microvascular blood flow in SCD. Their RBCs change shape upon de-oxygenation [5,31], and they display blood flow abnormalities including adhesion of RBCs, rolling and adhesion of leukocytes, and stasis in post-capillary venules [4,31–33]. Upon pathological analysis, there is tissue damage in multiple organs including the kidney, liver, lung, and spleen [34–36]. Like human SCD patients, transgenic bS mice display numerous signs of an inflammatory response such as elevated white counts and enhanced activation of NF-kB, a transcription factor critical for the inflammatory response . In addition, there are significant increases in the expression of endothelial cell adhesion molecules and the acute phase response protein serum amyloid P-component (SAP) . Sickle mice exhibit biochemical footprints consistent with excessive reactive oxygen species (ROS) generation, even at ambient air . These footprints are exaggerated further after exposure to hypoxia–reoxygenation resulting in enhanced activation of NF-kB and a pro-adhesive endothelial cell phenotype which is conducive to the formation of vascular obstructions [4,5]. We have incorporated the DSFC model  to observe blood flow changes in the subcutaneous tissue of normal and transgenic sickle mice. This DSFC model allows repeat visualizations of the same vessels over several hours, days, or even weeks and analysis of vessel diameters, RBC velocities, vascular stasis, and leukocyte rolling and adhesion after administering proinflammatory insults such as hypoxia–reoxygenation and TNF-a. The role of inflammation and the molecular events leading to the formation and resolution of vascular obstructions in SCD are still largely speculative. Because vaso-occlusion is a critical event in the pathology of SCD, we have used the DSFC and intravital microscopy to examine blood flow in the subcutaneous microcirculation of bS/S-Antilles mice and examined the effects of pro-inflammatory insults on leukocyte interactions with the endothelium and the genesis and resolution of vascular stasis. MATERIALS AND METHODS Mice All animal experiments were approved by the University of Minnesota’s Institutional Animal Care and Use Committee. We utilized female bS/S-Antilles transgenic mice as our mouse model for SCD . The bS/S-Antilles mice are homozygous for deletion of the mouse b major globin locus [bMDD] and express human a, bS, and bS-Antilles globin transgenes. These mice were produced by breeding aHbS[bMDD] mice  into a mouse line expressing human bS-Antilles on the mouse homozygous b major deletional background . bS-Antilles contains, in addition to the bS mutation at b6, a second mutation at b23 (Val!Ile). bS-Antilles has low oxygen affinity and decreased solubility under deoxygenated conditions, resulting in a more severe form of SCD. In the bS/S-Antilles mice, approximately 42% of the b globins expressed are human bS and 36% are human bS-Antilles. These transgenic bS/S-Antilles mice are on the C57BL/6 genetic background. Normal female mice (C57BL/6) obtained from Jackson Laboratory (Bar Harbor, ME) were used as normal controls for the bS/S-Antilles mice. The mice used in these studies were between 8 and 12 weeks of age. The mice weighed 20–30 g and were housed in specific pathogen-free housing to prevent common murine infections that could cause an inflammatory response. All the mice were maintained on a standard chow diet. Hematocrits, Reticulocytes, and White Counts Blood was collected from the tail vein of unanesthetized mice after warming the animals under a heat lamp. Heparinized whole blood was collected in capillary tubes for manual measurement of packed RBC volume (hematocrit) and for reticulocyte staining and counting in methylene blue. Reticulocytes, identified by their DNA staining, were expressed as a percentage of RBCs. For white counts, the RBCs were immediately lysed by diluting whole blood 20-fold in 2% acetic acid containing 30 mg/mL EDTA. Leukocytes were counted on a hemocytometer four times and averaged. Anesthetic Preparation Anesthesia was administered to all mice before surgery and intravital microscopy. The anesthetic cocktail, administered intraperitoneally, consisted of 0.2 mL of xylazine (20 mg/mL) and 0.6 mL of ketamine (100 mg/mL) mixed with 9 mL of normal saline. We injected 0.2–0.4 mL of this anesthetic cocktail into each mouse. All controlled substances used were approved by Research Animal Resources at the University of Minnesota. DSFC Implantation The experimental model used in this study was originally developed for the rat dorsal skin fold  and modified for mice. After the mouse was anesthetized, a suitable location for chamber placement was selected, Stasis in Sickle Cell Disease and the hair on that location was shaved off with a clipper. The remaining hair was removed from the dorsal skin with Nair Hair Remover (Church & Dwight, Princeton, NJ). The smooth skin was washed twice with betadine and dried. A sterile field was maintained throughout the surgical procedure. One sterilized chamber piece containing a round window was placed on one side of the dorsal skin fold. Another matching chamber piece without a window was placed on the opposite side of the skin fold, holes for three connecting screws were made through the skin fold and the assembly was tightened together with screws and nuts. A circle of cutaneous tissue and fascia was carefully cut away from the skin fold inside the DSFC window exposing the blood vessels of the subcutaneous tissue adjacent to the striated muscles of the opposing skin fold. Antibiotic ointment (bacitracin zinc/polymyxin B sulfate/neomycin sulfate) was used on the edges of the wound inside the DSFC window. A glass window was placed in the chamber to cover the exposed tissue and secured with a snap ring. Finally, the chamber was sutured to the skin. After surgery, the animals were housed in barrier cages inside a humidified incubator maintained at 32 C with unrestricted access to food and water. To minimize the risk of infection, all mice were given the antibiotic amoxicillin in their drinking water (250 mg per 5 mL of water) after DSFC implantation and throughout the experiment. The mice had a mild inflammatory response to the DSFC implantation, as evidenced by elevated levels of serum amyloid P component (SAP)  3 days after surgery (data not shown). SAP returned to pre-surgical baseline levels 4–7 days after DSFC implantation. Venule diameters, RBC velocity, vascular stasis, and leukocyte rolling and adhesion measurements, presented below, were measured on days 4–7 after DSFC implantation, and measurements in normal and sickle mice were made on the same days. Similar stasis and leukocyte rolling values were obtained on days 4–7 (data not shown) in both sickle and normal mice, indicating that the days chosen for data collection did not influence the results. Figure 1 shows a close-up of an implanted DSFC with the vasculature visible through the window. Intravital Microscopy and Analysis Four to seven days after DSFC implantation, anesthetized mice were placed on a specially constructed microscope stage where individual flowing venules were selected at random and their relative locations were noted with a hand drawn map of the microscopic field. Figure 2 shows a mouse mounted on the special microscope stage for intravital microscopy. The DSFC was illuminated with a 30-W halogen bulb. Microscopic observations were carried out in bright field using a Nikon Labophot-2 microscope (Fryer 119 Fig. 1. Close up of a dorsal skin fold chamber (DSFC) implanted on a transgenic sickle mouse. Subcutaneous blood vessels of the opposing dorsal skin fold are visible through the chamber window. Fig. 2. Mouse with an implanted DSFC mounted on the microscope stage prior to intravital microscopy of microvessels inside the DSFC window. Corp., Minneapolis, MN) fitted with a Hamamatsu Newvicon television camera (North Central Instruments, Minneapolis, MN) and a JVC S-VHS video recorder (JVC Company of America, Aurora, IL). The video images were subsequently digitized from the videotapes using ImagePro (Leeds Precision Instruments, Minneapolis, MN). Mean vessel diameters were measured offline using a digital image splitting device . Mean RBC velocities (VRBC centerline) were measured online using an optical Doppler velocimeter  obtained from Texas A&M University Microcirculation Research Institute. Wall shear rates were calculated using the formula 8Vmean/D, where D is the vessel diameter and Vmean is the mean velocity obtained from the equation Vmean ¼ VRBC centerline/1.6, as previously reported . Each vessel was observed online for blood flow. Venules with no observable blood flow were reported as static. 120 Kalambur et al. For measurement of leukocyte rolling and adhesion, the leukocytes were labeled in vivo with 100 mL of 0.02% rhodamine 6G (Sigma Chemical Co., St. Louis, MO) in sterile saline administered intravenously via tail vein injection . The DSFC was illuminated with a mercury lamp and rhodamine signal-enhancing filters (excitation l ¼ 480–520 nm and emission l ¼ 535– 585 nm) were used to view the fluorescence. A Hamamatsu silicon-intensified transmission camera (North Central Instruments) fitted to the Nikon Labophot-2 microscope was used to detect the fluorescent signal. Each vessel was recorded on videotape for approximately 2 min. Rolling leukocytes were measured offline using the videotape and were defined as those leukocytes that distinctly roll along the endothelial surface of venules . The rolling flux was determined as the total number of leukocytes rolling through a given section of vessel per minute. A leukocyte was considered adherent if it remained stationary for at least 30 sec. rank sum test. The proportions of venules exhibiting stasis at each time point were compared using a z-test. RESULTS The hematocrits (mean ± SE) of sickle (46.3 ± 3.0%, n ¼ 3) and normal (45.6 ± 0.8 %, n ¼ 7) mice were similar, although the number of reticulocytes in sickle mice (10.1 ± 1.0%, n ¼ 4), expressed as percent of RBCs, was 3.5-fold higher than normal mice (2.9 ± 0.7%, n ¼ 3, P < 0.01). Mean white counts in sickle mice ([14.3 ± 1.1] 103/mL, n ¼ 12) were 1.4-fold higher than normal mice ([8.7 ± 0.6] 103/mL, n ¼ 14, P < 0.001). A scatter plot of RBC velocities as a function of diameter within the venules (Fig. 3A) and the arterioles (Fig. 3B) of normal mice with implanted DSFCs at baseline ambient air conditions are compared with published values for the cat mesentery and rabbit Experimental Protocol Four to seven days after the DSFC implantations, anesthetized normal and bS/S-Antilles (sickle) mice were visualized separately and the venule diameters, mean RBC velocities, stasis, and leukocyte rolling and adhesion were measured as described above. After baseline measurements in ambient air, the mice were transferred to a special chamber and subjected to 1 hr of hypoxia (7% O2/93% N2) followed by reoxygenation in ambient/room air. Additional measurements of blood flow parameters were made at various times (0–24 hr) after hypoxia during the reoxygenation phase. Some anesthetized mice with DSFCs were injected intraperitoneally with recombinant mouse TNF-a (25 mg/kg body weight, R&D Systems, Minneapolis, MN) in ambient air. Vascular stasis and leukocyte rolling and adhesion were measured as described above 0–8 hr after TNF-a injection. Histology of Dorsal Skin In some sickle mice, dorsal skin samples were taken for histological analysis after 1 hr of hypoxia and 1 hr of reoxygenation. Skin samples were fixed overnight in formalin, cut into 5-mm sections, embedded in paraffin, mounted on slides, and stained with hematoxylin and eosin before microscopic examination. Statistics All statistical analyses were performed with SigmaStat 2.0 for Windows (SPSS Inc., Chicago, IL). Comparisons of data from normal and transgenic sickle mice in ambient air and after hypoxia–reoxygenation were made using Student’s t-test or a Mann–Whitney Fig. 3. Scatter plots of mean RBC velocities (mm/sec) versus mean diameters (mm) in venules (A) and arterioles (B) in normal mice with DSFCs compared to other animal model systems. Values were obtained from subcutaneous venules (n = 60) and arterioles (n = 42) in normal mice with implanted DSFCs. For comparison, data from the mesentery of cats (n = 48 venules and n = 44 arterioles) and the omentum of rabbits (n = 32 venules and n = 34 arterioles) were taken from the literature . Stasis in Sickle Cell Disease 121 TABLE I. Hypoxia–Reoxygenation Reduces Mean Diameters, RBC Velocities, and Wall Shear Rates in the Subcutaneous Venules of Sickle, But Not Normal, Mice Venule measurementsa Diameter (mm) RBC velocity (mm/sec) Wall shear rate (1/sec) Mouse model Number of venules (n) Baseline ambient air Normal Sickle Normal Sickle Normal Sickle 21 43 21 43 21 43 37.7 ± 4.0 33.2 ± 3.9 3.4 ± 0.3 4.2 ± 0.4 507 ± 50 979 ± 122B ! 1 hr hypoxia + 1 hr reoxygenation 37.3 ± 3.7 30.1 ± 3.8AA,B 4.3 ± 0.7 1.9 ± 0.3AAA,BBB 627 ± 88 450 ± 74AAA,B a Venule diameters, RBC velocities, and wall shear rates at baseline and after 1 hr of hypoxia and 1 hr of reoxygenation in normal and sickle mice. Values are presented as mean ± SE. AA P < 0.01 and AAAP < 0.001 when measurementstaken after hypoxia–reoxygenation are comparedtothose made at ambientair within the same venules. B P < 0.05 and BBBP < 0.001 when the measurements made in sickle mice are compared to those made in normal mice under the same treatment conditions. omentum . The RBC velocities in venules and arterioles of normal mice with DSFCs compare favorably with values of blood flow in cat mesentery and rabbit omentum model systems. The DSFC affords the advantage of examining the same venules at multiple time points before, during, and after a vaso-active intervention such as hypoxia–reoxygenation. Table I summarizes the blood flow measurements in normal and transgenic sickle mice in venules inside the subcutaneous DSFC microcirculation at ambient air baseline and after 1 hr of hypoxia and 1 hr of reoxygenation treatment. The mean venule diameter remained constant after hypoxia–reoxygenation in normal mice but decreased 9% (P < 0.01) in the sickle mice after hypoxia–reoxygenation. In the normal mice, there was a non-statistically significant 26% increase in the mean RBC velocity after hypoxia–reoxygenation, but in sickle mice the mean RBC velocity decreased 55% (P < 0.001) after hypoxia–reoxygenation. Similarly, the calculated wall shear rates increased an average of 24% in normal mice (not significant) after hypoxia–reoxygenation, but decreased 54% (P < 0.001) in sickle mice after hypoxia–reoxygenation. Baseline wall shear rates in sickle mice in ambient air were 93% greater than normal mice under the same conditions (P < 0.05). But, wall shear rates in sickle mice dropped precipitously to 72% of the wall shear rates in normal mice after hypoxia– reoxygenation (P < 0.05). We used the DSFC model to examine the genesis and resolution of microvascular stasis after hypoxia– reoxygenation in the venules of sickle and normal mice (Fig. 4). All of the selected venules were flowing at ambient air baseline (H,R ¼ 0,0) in normal (n ¼ 80) and sickle mice (n ¼ 178). None of the venules in normal mice became static at any time during hypoxia–reoxygenation, and none of the venules in sickle mice became static during the 1 hr of hypoxia or during the first 15 min of reoxygenation (H,R ¼ 1,0). However, after 1 hr of hypoxia and 1 hr of reoxygenation (H,R ¼ 1,1), 11.9% of the venules in sickle mice Fig. 4. Stasis occurs in the subcutaneous venules inside the DSFC of transgenic sickle mice, but not normal mice, after 1 hr of hypoxia and 1 hr of reoxygenation. Subcutaneous venules inside DSFCs implanted on normal and sickle mice were visualized by intravital microscopy. Flowing venules were randomly selected at ambient air baseline [H(ypoxia),R(eoxygenation) = 0,0]. After baseline examination of venules, the mice were exposed to 1 hr of hypoxia (7% oxygen and 93% nitrogen) and the same venules were re-examined for blood flow/stasis at the end of hypoxia during the first 10 min of reoxygenation (H,R = 1,0). Similarly, the same venules were re-examined for blood flow after 1 hr of hypoxia and 1 hr of reoxygenation (H,R = 1,1) and after 1 hr of hypoxia and 4 hr of reoxygenation (H,R = 1,4). Venules without any visible blood flow were counted as static and those with blood flow were counted as flowing. Each bar represents the percentage of static venules seen in the DSFC window. The total number of venules (n) examined was 178 in sickle mice and 80 in normal mice at each time point. **P < 0.001 and *P = 0.01 in sickle mice compared to the same venules at ambient air baseline. became static (P < 0.001 compared to baseline). After 1 hr of hypoxia and 4 hr of reoxygenation (H,R ¼ 1,4), most of the stasis had resolved; only 3.6% of the 122 Kalambur et al. TABLE II. Leukocyte Rolling and Adhesion in the Subcutaneous Venules of Normal and Sickle Mice at Ambient Air Baseline and After 1 hr of Hypoxia and 1 hr of Reoxygenation* Treatment Baseline at ambient air 1 hr hypoxia + 1 hr reoxygenation Mouse model Total number of venules (n) Leukocyte rolling flux (leukocytes/min) Leukocyte adhesion (leukocytes/100 mm) Normal Sickle Normal Sickle 17 16 13 25 2.5 + 0.6 13.4 ± 3.5BB 3.0 + 1.2 38.7 ± 6.1AA,BBB 0.7 + 0.3 2.6 ± 0.5BB 1.3 + 0.4 8.1 ± 1.0AAA,BBB *Values are presented as means ± SE. AA P < 0.01 and AAAP < 0.001 when leukocyte rolling and adhesion values after hypoxia–reoxygenation are compared to those at baseline ambient air within the same mice. BB P < 0.01 and BBBP < 0.001 when values in sickle mice are compared to those in normal mice under the same treatment conditions. subcutaneous venules in sickle mice remained static (P ¼ 0.01 compared to baseline, n ¼ 111 venules examined) and all of the venules were flowing again after 24 hr of reoxygenation (data not shown). The static venules in sickle mice had a smaller mean diameter than the nonstatic venules. The mean baseline diameter of the venules that became static in sickle mice was 39.0 ± 18 mm (mean ± SE), while the non-static venules had a mean baseline diameter of 47.5 ± 50.2 mm. TNF-a was injected intraperitoneally (25 mg/kg body weight) into sickle mice to see if the pro-inflammatory cytokine could induce stasis in a manner similar to hypoxia–reoxygenation treatment. None of the subcutaneous venules in sickle mice became static 0–8 hr after TNF-a injection despite significant increases in leukocyte rolling (data not shown). Leukocyte rolling flux and leukocyte firm adhesion were measured in the subcutaneous venules of normal and sickle mice at ambient air baseline and after 1 hr of hypoxia and 1 hr of reoxygenation (Table II). Leukocyte rolling flux and adhesion were significantly higher in the venules of sickle mice compared to those in normal mice at both ambient air baseline (P < 0.01) and after hypoxia–reoxygenation (P < 0.001). Hypoxia–reoxygenation increased leukocyte rolling flux (P < 0.01) and adhesion (P < 0.001) in transgenic sickle mice approximately 3-fold; however, in normal mice, hypoxia–reoxygenation had little effect on leukocyte rolling and adhesion. Rhodamine fluorescent staining of leukocytes revealed that there was an overabundance of firmly adherent leukocytes at sites of vascular stasis (Fig. 5A). Venules that become static are filled with adherent leukocytes at bifurcations near the ‘‘head’’ or beginning of the static venule (Fig. 5A). In data not shown, a video taken of the same venule in Fig. 5A shows that the venule is plugged with adherent leukocytes near a bifurcation that is continuous with a larger flowing venule. The leukocytes are labeled with rhodamine 6G and appear as white fluorescent balls. The video also shows some of the leukocytes become dislodged as the static vessel attempts to re-establish flow. Fig. 5. (A) Fluorescent image of a static venule with adherent leukocytes after hypoxia-reoxygenation in the DSFC implanted on a sickle mouse. Leukocytes were labeled in vivo with intravenous rhodamine 6G and appear as white balls in the figure. Adherent leukocytes can be seen at a bifurcation plugging the ‘‘head’’ or beginning of a static venule. The static venule is continuous with a larger flowing venule. The image was captured after 1 h of hypoxia (7% O2/93% N2) and 1 h of reoxygenation in room air. (B) Histology of venule in the dorsal skin of transgenic sickle mice after 1 hr of hypoxia and 1 hr of reoxygenation. Dorsal skin samples were taken for histological analysis after the sickle mice were exposed to 1 hr of hypoxia and 1 hr of reoxygenation when approximately 12% of the venules were static. Skin samples were fixed overnight in formalin, cut into 5-mm sections, embedded in paraffin, mounted on slides, and stained with hematoxylin and eosin before microscopic examination. The figure shows a venule with a suspected vascular obstruction. White arrowheads point to leukocytes that appear to be adherent to the vascular endothelium and white arrows point to misshapen RBCs inside the venule. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.] Stasis in Sickle Cell Disease Dorsal skin samples were taken for histological analysis from sickle mice after exposure to 1 hr of hypoxia and 1 hr of reoxygenation when approximately 12% of the subcutaneous venules become static. Figure 5B shows hematoxylin and eosin staining of a subcutaneous venule with a suspected vascular obstruction. White arrowheads point to leukocytes that appear to be adherent to the vascular endothelium, and white arrows point to misshapen RBCs inside the venule. DISCUSSION The development of transgenic murine models to study SCD provides an opportunity to understand the molecular and cellular events during the genesis and resolution of vaso-occlusion. Characterizing and understanding the molecular basis for vaso-occlusion in SCD is of critical importance to devising strategies to prevent and treat it. By its nature, vaso-occlusion and its subsequent resolution promote ischemia–reperfusion injury, which can induce ROS production, inflammation, and organ pathology. These studies utilized DSFCs in conjunction with intravital microscopy to examine the evolution of vascular stasis after hypoxia– reoxygenation in transgenic mice expressing human a and bS/S-Antilles globins. These studies show that vascular stasis, induced by hypoxia–reoxygenation, is a rapid process of relatively short duration; stasis occurs in the subcutaneous venules within 1 hr of reoxygenation and resolves largely by 4 hr and completely within 24 hr. The stasis is accompanied by a significant increase in leukocyte rolling and firm adhesion to the endothelium of venules with plugs of adherent leukocytes present at bifurcations at the ‘‘head’’ or beginning of the static venule continuous with larger flowing venules. Intravital microscopy of the subcutaneous microcirculation in the murine DSFC model provides blood flow measurements that are comparable to vascular beds in other animal models. The DSFC is advantageous to examine temporal changes in the same vessels, and the effects of interventions such as hypoxia–reoxygenation and TNF-a in transgenic bS mouse models. The DSFC allows animals to be observed over hours, days, or even weeks, without sacrificing the animal after several hours as is required in the cremasteric, mucosal–intestinal, and omentum preparations. In addition, unlike the cremasteric model, the DSFC model allows the study of the microcirculation in both male and female mice. Such sex comparisons are warranted given the reported disparities in nitric oxide production in males and females . The relationship of RBC velocities to venular and arterial diameters in normal mice with a DSFC was similar to the cat mesentery and rabbit omentum models as well as diameters and velocities in venules 123 of the mouse cremasteric model . Placing the chamber on the mice does cause a mild inflammatory response that subsides over time as the wound heals as indicated by SAP values (data not shown). A surgically induced inflammatory response is also a concern in the other intravital microscopy models as well. However, in the bS/S-Antilles and normal mice, the percentage of static venules and leukocyte rolling at baseline and after hypoxia–reoxygenation were similar on days 4–7 after DSFC implantation (data not shown). This indicates that any inflammatory response that may have been present on days 4–7 after DSFC surgery did not have a significant impact on stasis and rolling in the subcutaneous venules. In our studies, using a bS/S-Antilles murine model with generally moderate severity of SCD , differences in blood flow were observed in subcutaneous venules when sickle mice were compared to normal mice. Of note, at ambient air baseline, the sickle mice had significantly faster blood flow and higher wall shear rates than normal mice. This finding contrasts with reports of slower blood flow in bS/S-Antilles mice in ambient air in the post-capillary venules of cremasteric muscle preparations . This difference could be due to differences in the vascular beds or perhaps to differences in the level of baseline vascular inflammation between the two models. In preliminary studies in our laboratory, adaptive, cytoprotective genes such as heme oxygenase1 are induced in SCD and enzymatic products from heme destruction such as carbon monoxide and biliverdin have vasodilatory and anti-inflammatory properties that may compensate and protect the vascular endothelium and alter blood flow in the absence of an inflammatory stress such as hypoxia. After hypoxia– reoxygenation, a pro-inflammatory insult, the venules of sickle mice with DSFCs had markedly slower blood flow and lower wall shear rates than normal mice; a similar finding was reported for the cremasteric muscle preparation . Unlike normal mice, sickle mice were extremely sensitive to hypoxia–reoxygenation. Dramatic reductions in blood flow in response to hypoxia–reoxygenation were accompanied by 3-fold increases in leukocyte rolling and firm adhesion. In addition to increased leukocyte rolling and adhesion, sickle mice also responded to hypoxia– reoxygenation treatment with the development of stasis in the subcutaneous venules inside the DSFC after 1 hr of reoxygenation. The stasis in sickle mice was significantly reduced after 4 hr of reoxygenation and completely absent after 24 hr of reoxygenation. There were no signs of stasis in any normal animals. Histological analysis of the dorsal skin from sickle mice after hypoxia–reoxygenation revealed leukocytes and misshapen RBCs that appear to be adherent to the endothelium in suspected static venules. 124 Kalambur et al. Previous studies of the blood flow dynamics in bS mice were performed in the mucosal–intestinal  and cremasteric [4,31,33] vascular beds. Intravital microscopy studies of blood flow in the cremasteric muscle preparation of bS mice showed adhesion of RBCs and leukocytes in the post-capillary venules , and studies in the mucosal–intestinal microcirculation detected sludging and decreased blood flow velocities in venules of all diameters in bS mice compared to normal . Some of these studies induced inflammation in the cremasteric venules by exposing the animals to hypoxia–reoxygenation  or by administration of the pro-inflammatory cytokine TNF-a . Hypoxia– reoxygenation increased endothelial ROS production, leukocyte rolling flux and leukocyte firm adhesion and decreased leukocyte rolling velocities and venular blood flow in bS mice compared to normal . In severe, but not milder, mouse models of SCD, TNF-a induced stasis in the cremasteric venules and was fatal in a high percentage of mice transplanted with bone marrow from the severe bS ‘‘BERK’’ mouse models . Inhibition of P-selectin binding, but not E-selectin, in bS mice during hypoxia–reoxygenation essentially eliminated leukocyte rolling and firm adhesion and markedly increased RBC velocities and wall shear rates . Similarly, mice deficient in P- and E-selectin, transplanted with bone marrow from ‘‘BERK’’ bS mice, had reduced leukocyte rolling and adhesion and were protected from TNF-a induced stasis . Thus, these intravital microscopy studies of blood flow in inflamed subcutaneous and cremasteric venules of bS and bS+S-Antilles mice and normal mice transplanted with bone marrow from bS mice suggest that endothelial cell activation and the accompanying leukocyte rolling and adhesion may be linked to vascular stasis in SCD. However, because we could not induce stasis with the pro-inflammatory cytokine TNF-a, despite increases in leukocyte rolling and adhesion seen 4 hr after TNF-a administration, factors other than adhesion molecule expression may be involved in stasis. These data suggest that sickling, and perhaps hemolysis of the red cell, which occurs during hypoxia, may be required before stasis can occur; TNF-a induced inflammation, alone, is not sufficient to induce stasis in less severe models of SCD such as the bS+S-Antilles. CONCLUSIONS In summary, the DSFC model provides an excellent tool for examining vascular stasis, RBC velocity, and leukocyte interactions with the endothelium in the subcutaneous microcirculation of transgenic sickle mice. This model may be relevant in SCD as the subcutaneous vasculature is frequently occluded and may be the primary cause of skin ulcerations in humans with SCD . The development of stasis and its resolution, as seen in the DSFC, is by its nature consistent with a model of on-going ischemia–reperfusion injury in transgenic sickle mice and may be perhaps fundamental to the pathology of SCD. Vascular inflammation could lead to a vicious cycle of inflammation and stasis, where inflammation leads to leukocyte and RBC adherence to endothelium, followed by stasis and temporary local ischemia, resolution, and reperfusion leading to ROS production, more inflammation, adhesion, and stasis. The DSFC model in transgenic sickle mice affords a potential avenue for discovery of the molecular and cellular events leading to stasis and its resolution as well as the testing of potential therapies. The model needs to be tested on other transgenic sickle animals that have varying expression of human g globins in conjunction with the expression of human a and bS globins and varying severity of SCD [48,49]. This model can also be used to examine the role of fluorescently labeled sickle RBCs in the formation of vascular obstructions. We hope that the insights derived from this model will be readily translated to humans with SCD to improve blood flow, minimize vasoocclusion, and prevent organ injury. ACKNOWLEDGMENTS We thank Dr. Kenneth Diller, Department of Mechanical Engineering, University of Texas, Austin, Dr. Marcos Intaglietta, Department of Bioengineering, University of California, San Diego, and their colleagues for their assistance on intravital microscopy setup and enhancement. We also thank Stephana Choong for breeding and characterizing the transgenic sickle mice used for these studies and Julia Nguyen for her help with blood collection. REFERENCES 1. Embury SH, Hebbel RP, Steinberg MH, Mohandas N. Pathogenesis of vasoocclusion. In: Embury SH, Hebbel RP, Mohandas N, Steinberg MH, editors. Sickle cell disease: basic principles and clinical practice. New York: Raven Press; 1994. Vol 21, p 311–326. 2. Pauling L, Itano HA, Singer SJ. Sickle cell anemia, a molecular disease. Science 1949;100:543–548. 3. Bunn FH. Pathogenesis and treatment of sickle cell disease. N Engl J Med 1997;337:762–769. 4. Kaul DK, Hebbel RP. Hypoxia/re-oxygenation causes inflammatory response in transgenic sickle mice but not in normal mice. J Clin Invest 2000;106:411–420. 5. Osarogiagbon UR, Choong S, Belcher JD, Vercellotti GM, Paller MS, Hebbel RP. Reperfusion injury pathophysiology in sickle transgenic mice. Blood 2000;96:314–320. 6. Buchanan GR, Bowman WP, Smith SJ. Recurrent cerebral ischemia during hypertransfusion therapy in sickle cell anemia. J Pediatr 1983;103:921–923. Stasis in Sickle Cell Disease 7. Castro O, Brambilla DJ, Thorington B, et al. The acute chest syndrome in sickle cell disease: incidence and risk factors. The Cooperative Study of Sickle Cell Disease. Blood 1994;84:643–649. 8. Vichinsky EP, Styles LA, Colangelo LH, Wright EC, Castro O, Nickerson B. Acute chest syndrome in sickle cell disease: clinical presentation and course. Cooperative Study of Sickle Cell Disease. Blood 1997;89:1787–1792. 9. Amogu AU. Leucocyte counts in children with sickle cell anemia usefulness of stable state values during infections. West Afr J Med 2000;19:55–58. 10. Hedo CC, Aken’ova YA, Okpala IE, Durojaiye AO, Salimonu LS. Acute phase reactants and severity of homozygous sickle cell disease. J Intern Med 1993;233:467–470. 11. Singhal A, Doherty JF, Raynes JG, et al. Is there an acute-phase reactant response in steady-state sickle cell disease? Lancet 1993; 341:651–653. 12. Stuart J, Stone PC, Akinola NO, Gallimore JR, Pepys MB. Monitoring the acute phase response to vaso-occlusive crisis in sickle cell disease. J Clin Pathol 1994;47:166–169. 13. Belcher JD, Marker PH, Weber JP, Hebbel RP, Vercellotti GM. Activated monocytes in sickle cell disease: potential role in the activation of vascular endothelium and vaso-occlusion. Blood 2000; 96:2451–2459. 14. Francis RB Jr, Haywood LJ. Elevated immunoreactive tumor necrosis factor and interleukin-1 in sickle cell disease. J Natl Med Assoc 1992;84:611–615. 15. Malave I, Perdomo Y, Escalona E, et al. Levels of tumor necrosis factor/cachetin (TNF alpha) in sera from patients with sickle cell disease. Acta Haematol 1993;90:172–176. 16. Croizat H. Circulating cytokines in sickle cell patients during steady state. Br J Haematol 1994;87:592–597. 17. Kuvibidila S, Gardner R, Ode D, Yu L, Lane G, Warrier RP. Tumor necrosis factor alpha in children with sickle cell disease in stable condition. J Natl Med Assoc 1997;89:609–615. 18. Wun T, Cordoba M, Rangaswami A, Cheung AW, Paglieroni T. Activated monocytes and platelet–monocyte aggregates in patients with sickle cell disease. Clin Lab Haematol 2002;24:81–88. 19. Hofstra TC, Kalra VK, Meiselman HJ, Coates TD. Sickle erythrocytes adhere to polymorphonuclear neutrophils and activate the neutrophil respiratory burst. Blood 1996;87:4440–4447. 20. Fadlon E, Vordermeier S, Pearson TC, et al. Blood polymorphonuclear leukocytes from the majority of sickle cell patients in the crisis phase of the disease show enhanced adhesion to vascular endothelium and increased expression of CD64. Blood 1998;91:266–274. 21. Lard LR, Mul FP, de Haas M, Roos D, Duits AJ. Neutrophil activation in sickle cell disease. J Leukoc Biol 1999;66:411–415. 22. Kenny MW, George AJ, Stuart J. Platelet hyperactivity in sicklecell disease: a consequence of hyposplenism. J Clin Pathol 1980; 33:622–625. 23. Westwick J, Watson-Williams EJ, Krishnamurthi S, et al. Platelet activation during steady state sickle cell disease. J Med 1983;14:17–36. 24. Beurling-Harbury C, Schade SG. Platelet activation during pain crisis in sickle cell anemia patients. Am J Hematol 1989;31: 237–241. 25. Papadimitriou CA, Travlou A, Kalos A, Douratsos D, Lali P. Study of platelet function in patients with sickle cell anemia during steady state and vaso-occlusive crisis. Acta Haematol 1993;89: 180–183. 26. Wun T, Paglieroni T, Tablin F, Welborn J, Nelson K, Cheung A. Platelet activation and platelet–erythrocyte aggregates in patients with sickle cell anemia. J Lab Clin Med 1997;129:507–516. 27. Wun T, Paglieroni T, Field CL, et al. Platelet–erythrocyte adhesion in sickle cell disease. J Invest Med 1999;47:121–127. 28. Inwald DP, Kirkham FJ, Peters MJ, et al. Platelet and leucocyte activation in childhood sickle cell disease: association with nocturnal hypoxaemia. Br J Haematol 2000;111:474–481. 125 29. Solovey A, Lin Y, Browne P, Choong S, Wayner E, Hebbel RP. Circulating activated endothelial cells in sickle cell anemia. N Engl J Med 1997;337:1584–1590. 30. Solovey A, Gui L, Key NS, Hebbel RP. Tissue factor expression by endothelial cells in sickle cell anemia. J Clin Invest 1998;101: 1899–1904. 31. Kaul DK, Fabry ME, Costantini F, Rubin EM, Nagel RL. In vivo demonstration of red cell–endothelial interaction, sickling and altered microvascular response to oxygen in the sickle transgenic mouse. J Clin Invest 1995;96:2845–2853. 32. Embury SH, Mohandas N, Paszty C, Cooper P, Cheung AT. In vivo blood flow abnormalities in the transgenic knockout sickle cell mouse. J Clin Invest 1999;103:915–920. 33. Turhan A, Weiss LA, Mohandas N, Coller BS, Frenette PS. Primary role for adherent leukocytes in sickle cell vascular occlusion: a new paradigm. Proc Natl Acad Sci USA 2002;99:3047–3051. 34. Fabry ME, Costantini F, Pachnis A, et al. High expression of human beta S- and alpha-globins in transgenic mice: erythrocyte abnormalities, organ damage, and the effect of hypoxia. Proc Natl Acad Sci USA 1992;89:12150–12159. 35. Rubin EM, Witkowska HE, Sprangler E, et al. Hypoxia-induced in vivo sickling of transgenic mouse red cells. J Clin Invest 1991; 87:639–647. 36. Fabry ME, Sengupta A, Suzuka SM, et al. A second-generation transgenic mouse model expressing both hemoglobin S (HbS) and HbS-Antilles results in increased phenotypic severity. Blood 1995; 86:2419–2428. 37. Belcher JD, Bryant CJ, Nguyen J, et al. Transgenic sickle mice have vascular inflammation. Blood 2003;101:3953–3959. 38. Papenfuss HD, Gross JF, Intaglietta M, Tresse FA. A transparent access chamber for the rat dorsal skin fold. Microvasc Res 1979; 18:311–318. 39. Fabry ME, Nagel RL, Pachnis A, Suzuka SM, Costantini F. High expression of human beta S- and alpha-globins in transgenic mice: hemoglobin composition and hematological consequences. Proc Natl Acad Sci USA. 1992;89:12150–12154. 40. Burlingame RW, Volzer MA, Harris J, Du Clos TW. The effect of acute phase proteins on clearance of chromatin from the circulation of normal mice. J Immunol 1996;156:4783–4788. 41. Hogan RD, Morris RF, McMurray SK. A digital video image splitting device for microvascular measurements. Microvasc Res 1984;27:128–132. 42. Borders JL, Granger HJ. An optical Doppler velocimeter. Microvasc Res 1984;27:117–127. 43. Davis MJ. Determination of volumetric flow in capillary tubes using an optical Doppler velocimeter. Microvasc Res 1987;34: 223–230. 44. Baatz H, Steinbauer M, Harris AG, Krombach F. Kinetics of white blood cell staining by intravascular administration of rhodamine 6G. Int J Microcirc Clin Exp 1995;15:85–91. 45. Zweifach BW, Lipowsky HH. Quantitative studies of microcirculatory structure and function. III. Microvascular hemodynamics of cat mesentery and rabbit omentum. Circ Res 1977;41(3):380–390. 46. Gladwin MT, Schechter AN, Ognibene FP, et al. Divergent nitric oxide bioavailability in men and women with sickle cell disease. Circulation 2003;107:271–278. 47. Koshy M, Entsuah R, Koranda A, et al. Leg ulcers in patients with sickle cell disease. WBC rolling flux and adhesion were significantly higher in the venules of sickle mice compared to those in normal mice at both ambient air baseline (P < 0.01) and after hypoxia–reoxygenation (P < 0.001). Blood 1989;74:1403–1408. 48. Paszty C, Brion CM, Manci E, et al. Transgenic knockout mice with exclusively human sickle hemoglobin and sickle cell disease. Science 1997;278:876–878. 49. Fabry ME, Suzuka SM, Weinberg RS, et al. Second generation of knockout sickle mice: the effect of HbF. Blood 2001;97:410–418.