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Expression of the cell adhesion molecule, L-selectin, on polymorphonuclear leukocytes during and after… Van Eeden, Stephanus Frederick 1994

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EXPRESSION OF THE CELL ADHESION MOLECULE, L-SELECTIN, ONPOLYMORPHONUCLEAR LEUKOCYTES DURING AND AFTER THEIRRELEASE FROM THE BONE MARROWbySTEPHANUS FREDERICK VAN EEDENMB.ChB. The University of Stellenbosch, South Africa, 1975M.Med. The University of Stellenbosch, South Africa, 1984FCP. The College of Physicians, South Africa, 1984A THESIS SUBMITTED IN PARTIAL FULFILMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIES(Department of Experimental Medicine)We accept this thesis as conformingto the r uired standard‘ZfrTHE UMVERSfl* OF BRItISH COLUMBIAAugust 1994©Stephanus Frederick van Eeden, 1994In presenting this thesis in partial fulfilmentof the requirements for an advanceddegree at the University of British Columbia,I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives, It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.Department of 4!,S4j%A’ eThe University of British ColumbiaVancouver, CanadaDate__________DE-6 (2/88)UABSTRACTThe emigration of polymorphonuclear leukocytes (PMN) to sites of inflammationrequires a series of leukocyte-endothelial interactions which can be divided in foursequential steps. These include, tethering of flowing leukocytes; triggering oractivation of leukocytes; firm adhesion to the endothelium and; emigration ofleukocytes out of the vessels into inflamed tissue. The selectin family of adhesionmolecules are involved in the tethering of leukocytes and one of these, L-selectin, isexpressed on nearly all leukocytes including PMN. The recruitment of the PMN tosites of inflammation is highly dependent on the expression of this molecule on theircell surfaces. However, the role of L-selectin in the trafficking of the PMN from thebone marrow into the blood is less clear. The objective of this thesis was todetermine the expression of L-selectin on PMN during their release from the bonemarrow into the circulation and their eventual removal into the tissues.Immunocytochemical and immunohistochemical techniques were used to determinethe expression of L-selectin on the PMN in cytological and histological specimensrespectively. Indirect immunofluorescent flow cytometry was used to determine theexpression of L-selectin on circulating PMN. Changes in L-selectin expression onPMN during their release from the bone marrow was studied during cardiopulmonarybypass in humans. These studies demonstrated that the expression of L-selectin onthe mature segmented PMN in bone marrow is higher than on circulating PMN andthat some of this L-selectin shed when PMN cross from the bone marrowmhematopoietic compartment into the bone marrow venous sinusoids. However, Lselectin expression on PMN in the venous sinusoids remains high and this results inan increased expression of L-selectin on circulating PMN during active bone marrowrelease. This means that circulating PMN expressing the highest levels of L-selectin,have been recently released from the bone marrow. A new method for labelling PMNin vivo using the thymidine analogue, 5’bromo-2-deoxyuridine was used todemonstrate that circulating PMN continuously lose L-selectin while they remain inthe circulation.These studies explain the variable expression of L-selectin on the circulating PMNwith the newly released PMN expressing the highest and the older PMN the lowestlevels of L-selectin. This implies that the expression of L-selectin on circulating PMNis not just a marker of cell activation state, but also of cell age. This creates theopportunity to study the functional capabilities and the trafficking of differentpopulations of the circulating PMN.lvTABLE OF CONTENTSAbstract iiTable of Contents ivList of Tables viList of Figures viiList of Abbreviations XAcknowledgements xiiiDedication xiv1) General Introduction 1a) Distribution of PMN in the bone marrow. 3b) Distribution of PMN in the vascular space. 7c) Distribution of PMN in the tissue. 102) Bone marrow release of PMN (review) 12a) The structural-functional relationship of the bonemarrow-blood barrier. 13b) Developmental changes and leukocyte egress from the bone marrow. 17c) Factors that cause release of PMN from the bone marrow. 19d) Adhesion events and egress of PMN from the bone marrow. 203) Selectins 27L-selectin 281) Structure of the L-selectin protein 282) Expression of L-selectin on leukocytes 323) Regulation of L-selectin expression 334) Ligands for L-selectin 355) Functions of L-selectin 364) L-selectin expression during active bone marrow release of PMNIntroduction 39Working hypothesis 42Specific Aims 42VModel of bone marrow release of PMN 43Methods 45Results 62Discussion 825) Changes in L-selectin on circulating PMNIntroduction 94Hypothesis 97Specific Aims 97Model of in vivo labelling of PMN 98Methods 101Results 111Discussion 122L-selectin expression on PMN during their lifespanin the circulation 127Methods 130Results 139Discussion 1476) Summary and future directions 1547) References 159viLIST OF TABLESTable I. The selectin group of cell adhesion molecules: Nomenclature,expression and function (p-29).Table 11. Clinical characteristics of cardiopulmonary bypass patients (p-63).Table III. Changes in the peripheral leukocyte counts in patients duringcardiopulmonary bypass (p-64).Table IV. Changes in the L-selectin expression on circulating PMN in patientsduring cardiopulmonary bypass (p-69).Table V. Changes in L-selectin expression on PMN in the bone marrow inpatients during cardiopulmonary bypass (p-Ti).Table VI. Comparison of the number of white cells, PMN and BrdU labelledPMN recovered in three different preparations of blood from rabbitslabelled in vivo with BrdU (p-i 13).Table VII. Half-lives of BrdU labelled PMN infused as either whole blood,leukocyte rich plasma or purified PMN (p-i 17).Table Vifi. Half-lives of BrdU labelled PMN treated with and withoutChymotrypsin infused into recipient (p-146).vuLIST OF FIGURESFigure 1: The stages of maturation and flux of polymorphonuclear leukocytes in thebone marrow, circulating blood and tissue compartments (p-5).Figure 2: Schematic presentation of the bone marrow-blood barrier (p-l5).Figure 3: Domain compositions of the three known selectins (p-31).Figure 4: Immunocytochemical grading system used to evaluate the intensity ofstaining of PMN in the circulation (p-55).Figure 5: The disector method used to quantify the number of PMN in each bonemarrow compartment (p-57).FIgure 6: Photomicrograph of a bone marrow sinusoid demonstrated with a ToluidineBlue 0 stain and the corresponding serial section stained for the presence of Lselectin (p-58).Figure 7: Variability in grading the expression of L-selectin on PMN in cytologicalspecimens (p-66).Figure 8: The correlation between the visual and Infrascan® grading systems (p-67).Figure 9: The difference between L-selectin expression on PMN in the circulatingblood (cytospins) and the bone marrow (bone marrow smears) (p-68).Figure 10: Immunocytochemical determined L-selectin expression on circulatingPMN at the start and the end of 5 normothermic and 5 hypothermic cardiopulmonarybypass procedures (p-7O).Figure 11: The expression of L-selectin on segmented PMN and band cells in thebone marrow and the circulating blood (p-72).Figure 12: Expression of L-selectin on PMN during bone marrow release of PMN ina normothermic and hypothermic patient as measured by flow cytometry (p-73).Figure 13: Expression of L-selectin on circulating PMN as measured by flowcytometry at the different time intervals in 5 normothermic and 5 hypothermic CPBprocedures (p-74).Figure 14: Immunohistochemical determination of L-selectin on PMN in bonemarrow hematopoietic and sinusoid compartments at baseline. Combined baselinedata for both normothermic and hypothermic groups (p-76).vu’Figure 15: Changes in the number of L-selectin negative graded PMN in the bonemarrow sinusoids at the start (BM1) and at the end (BM2) of CPB bypass (p-78).Figure 16: Flow cytometric data concerning the expression of L-selectin and CD18on PMN incubated at 37, 27 and 4°C in vitro and stimulated with increasingconcentrations of zymosan activated plasma ranging from 0 to 5 % (p-79).Figure 17: Immunocytochemical changes in the L-selectin negative graded PMNincubated at 37, 27 and 4°C stimulated with increasing concentrations of zymosanactivated plasma (p-81).Figure 18: Time course of BrdU incorporation into circulating PMN donor rabbitsinfused with 25 mg/kg/day of BrdU for 7 days (p-112).Figure 19: Immunocyto- histochemical detection of BrdU labelled cells in thecirculating blood, spleen and pneumonic lung of recipient rabbits (p-i 14).Figure 20: BrdU labelled PMN in recipient’s circulation as a fraction of the numberof BrdU labelled PMN infused after the infusion of whole blood, LRP or purifiedPMN (p-li5).Figure 21: Semi-quantitation of BrdU labelled DNA in recipient organs 24 h afterthe transfusion of whole blood and LRP (p-i 19).Figure 22: Immunological detection of BrdU labelled in DNA extracted from organsof one of the recipient rabbits after the transfusion of whole blood from a donor rabbit(p- 120).Figure 23: Immunocytochemical detection of BrdU labelled PMN in cytospinprepared from LRP of the donor blood. Double immunolabelling of PMN on cytospinsfor both nuclear BrdU (blue) and surface L-selectin (red) using a double alkalinephosphatase technique (p-i37).Figure 24: L-selectin changes on PMN during time spent in the circulation whenBrdU labelled PMN were infused as whole blood or purified PMN (p-i40).Figure 25: The effect of chymotrypsin on the expression of L-selectin and CD18 asmeasured with flow cytometry (p-141).Figure 26: BrdU labelled PMN in the circulation of recipients after the infusion ofpurified PMN treated with either denatured chymotrypsin or chymotrypsin (p-i43).Figure 27: BrdU labelled L-selectin positive and negative PMN in the circulation ofrecipients after the infusion of purified PMN treated with either denaturedchymotrypsin or chymotrypsin (p-i44).IxFigure 28: Changes in the expression of L-selectin on PMN during their normallifespan (p-i55)xLIST OF ABBREVIATIONSMicrogram(s)1il Microliter(s)60.3 Anti-CD18 MoAbACCR Adventitial cell covering rateAPAAP Alkaline phosphate anti-alkaline phosphateARDS Adult Respiratory Distress SyndromeBM Bone marrowBSA Bovine serum albuminBrdU 5 ‘bromo-2-deoxyuridineCD Cluster of differentiation designationCR Complement regulatory domainCAM Cell adhesion moleculeCPB Cardiopulmonary bypassDREG-200 Anti-L-selectin MoAbECM Extracellular matrixEGF Epidermal growth factor domainE-selectin Endothelial-selecting GramG0-G4 Grades 0-4 (see grading system figure 4)G-CSF Granulocyte colony stimulating factorGM-CSF Granulocyte monocyte colony stimulating factorxiGMA Glycolmethacrylateh Hour(s)ICAM Intercellular adhesion moleculekg KilogramLPS LipopolysaccharideLRP Leukocyte rich plasmaL-selectin Leukocyte-selectinMFI Mean fluorescence intensityMoAb Monoclonal antibodiesmg milligram(s)mm minute(s)P-selectin Platelet-selectin (also on endothelial cells)PBS Phosphate buffered salinePMN Polymorphonuclear leukocyteRGD Arginine-glycine-aspartic acid attachment peptides second(s)SD Standard deviationSE Standard error of the meansLex Sialylated Lewis x antigensL-selectin Soluble L-selectinTBS Tris buffered salineTBO Toluidine BluexliVCAM Vascular cell adhesion moleculeZAP Zymosan activated plasmaxfflACKNOWLEDGEMENTSI wish to acknowledge my supervisor, Dr James Hogg, for supporting me in my questto do a Ph.D. in the Pulmonary Research Laboratory and stimulating my interest inleukocyte kinetics. I am fortunate to have trained under James and I consider hima academic mentor and friend of outstanding value. Special thanks are extended tomy committee members Dr’s Keith Walley, Graham Dougherty and Donna Hogge fortheir patience and constructive criticism. I also want to thank Jenny Hards forsharing with me her time and knowledge on immunocyto- and histochemistry. I willalways be grateful for her kindness and friendship. To Dr’s Alan Burns, ClaireDoerschuk and Blaire Walker I owe a special debt of gratitude for many fruitfuldiscussion on adhesion molecules. The animal experiments were greatly facilitatedby the surgical expertise of Dean English and I am in debt to Simon Bicknell for hishelp in the animal experiments. Dr Robert Miyagashima and all the operatingtheatre staff deserves a special acknowledgement for their willingness and supportin collecting the bone marrow specimens. I would like to acknowledge the assistanceof Dr Lawrence Haley and the technical staff in Immunology for helping with theflow cytometry. I would also like to acknowledge the help of the technical staff of thePRL, Harvey Coxson (stereology), Lorri Verburgt and Barry Wiggs (statisticalanalysis), Joe Comeau (computers) and the always available Steward Greene(photography). Finally, I am grateful to Prof’s James Joubert and Attie de Kock forencouraging me to pursue a academic career and I would like to acknowledge thefinancial support I received from the Medical Research Council of South Africa,Faculty of Medicine University of Stellenbosch, Dancun Flockhard, fellowship fromUniversity of British Columbia and B.C. Lung Association.xivDEDICATIONTo my wife, Christelle, and my family for their continuous support that has madethis work a joy to complete.11) GENERAL INTRODUCTIONThe control of the number of circulating leukocytes has been a topic of interest andcontroversy for almost a century. This is especially true for polymorphonuclearleukocytes (PMN) because they are implicated in the pathogenesis of tissue injury inseveral diseases (Cochrane et aL, 1965; Hammerschmidt et al., 1980a; Kniker andCochrane, 1965; Phelps and McCarthy, 1966; Stetson, 1951; Winn et aL, 1973).Neutrophilic leukocytosis is a feature common to many types of infectious andinflammatory diseases and serves to control the expansion and dissemination ofmicrobial or viral infection by killing or removing the pathogenic organisms. In otherclinical settings such as in adult respiratory distress syndrome (ARDS) (Brigham etal., 1979; Hammerschmidt et at., 1980a; Heflin and Brigham, 1981; Meguire et al.,1982; Tate and Repine, 1983a), sepsis syndrome resulting in multi-organ failure,ischemia-reperfusion tissue injury, gouty arthritis (Phelps and McCarthy, 1966) andchronic tissue injury in pulmonary emphysema (Janoff, 1983), PMN are thought tocontribute to the extent of tissue damage. For example, complement-inducedgranulocyte aggregation and intravascular cell activation are consideredinstrumental in the pathophysiology of ARDS (Tate and Repine, 1983b; Rinaldo,1986). Entrapment of activated PMN in the lung (Weiland et al., 1986) perpetuatesa number of interlocking inflammatory pathways including the coagulation andfibrinolysis cascade, kinin system, complement pathways, and arachidonic acidmetabolism (Rinaldo, 1986; Hammerschmidt et al., 198Gb; Tracey et al., 1988).Activated PMN damage the pulmonary endothelium: 1) directly; 2) by the release ofproteases (elastase and collagenase); 3) by the simultaneous activation of coagulationand fibrinolysis; 4) by the generation of arachidonic acid metabolites; 5) and by the2release of oxygen-free radicals (Brigham, 1977; Brigham and Owen, 1975; Cochraneet al., 1983; Rinaldo, 1986; Tate and Repine, 1983b). Evidence for PMN participatingin the development of ARDS is substantial (Tate and Repine, 1983b), and PMNdepletion has been shown to prevent ARDS-like injury after the injection of PMNactivators in a variety of animal models (Heffin and Brigham, 1981). Although ARDShas been described in neutropenic patients, PMN reconstitution worsens the ARDSthat develop in these patients (Rinaldo and Borovetz, 1985). Furthermore, thepathology in tobacco smoke related lung disease is located in the peripheral airways(Hogg et al., 1968) and is thought to be due to an inflammatory process involvingPMN (Wright et aL, 1988; Bosken et al., 1992) initiated by the cigarette smoke(Neiwoehner et al., 1974). Baseline circulating leukocyte counts have been shown tobe elevated in chronic smokers (Corre et al., 1971) and these PMN passing throughthe pulmonary capillary bed are exposed to components of cigarette smoke that delay(Bosken et al., 1992) and activate them (Klute et al., 1993) which could releaseproteases and reactive oxygen intermediates that damage the pulmonary endotheliumfrom within the vascular space. Regulation of the circulating PMN levels is thereforeimportant in mounting a response to bacterial infections and in controlling ormaintaining other inflammatory states.The total number of PMN in the body can be broken down into those in the bonemarrow, the circulating, the marginated and tissue pools. The number of PMN thatcirculate can be influenced by several factors including: the rate of production in themarrow; the rate of release from the marrow; the exchange between the circulatingand marginated pool of intravascular PMN; and their rate of permanent removal3from the circulation into the tissues. The PMN that enter the circulation duringstressful stimuli come from either a pool of cells marginated along the vessel wallswithin the vascular space and/or from the bone marrow (Doerschuk and Allard, 1989;Muir et aL, 1984). Mobilization of PMN from the marginated pool does not effectivelyincrease the total number of PMN in the vascular system. Alternatively, the releaseof PMN from the bone marrow increases the total number of PMN in the circulatingand marginated poois and therefore in cells that may participate in the inflammatoryresponse.a) Distribution of PMN in the bone marrowPolymorphonuclear leukocytes are produced in the bone marrow which is ahematopoietic factory weighing approximately 2600g, or 4.5 % of the body weight ofa normal adult. The bone marrow is widely distributed throughout the skeleton andas a single organ is larger than the liver (which weighs ± 1500g). About 55 %-60 % ofthe bone marrow is dedicated to the production of a single cell type, the neutrophil,with a normal ratio of neutrophils to erythrocytes of 2:1 to 3:1. Other granulocytescontribute about 3 %-5 % of the bone marrow. The cellularity of the bone marrowvaries with age and accounts for 75 % of the marrow in the young, 50 % in youngadults and as low as 25 % in the elderly (Hartsock et al., 1965). There is, however, agreat variability between individuals of any given age. Three major compartmentsof differentiation in the bone marrow have been distinguished: 1) the most primitivecompartment of pluripotent stem cells; 2) committed cells and; 3) maturing cells. Innormal adults, the life of PMN is spent in three environments- the bone marrow,blood and tissue. Bone marrow is the site of proliferation, terminal differentiation4and maturation of neutrophilic granulocytes. Proliferation consists of approximately5 divisions and only takes place during the first three stages of PMN development(myeloblast, promyelocyte and myelocyte stages). After the myelocyte stage the cellsbecome “end cells” that are no longer capable of dividing (Bainton et aL, 1971). Theythen enter a large storage pool where they mature to segmented PMN for ±5 daysbefore being released into the blood (Bainton et at., 1971; Bainton, 1980). Figure 1illustrates the PMN lifespan and stages of maturation (Bainton et aL, 1971; Bainton,1980; Wintrobe, 1981). Of every 100 nucleated cells in the bone marrow, 2% aremyeloblasts, 5 % promyelocytes, 12 % myelocytes, 20% metamyelocytes and band cells,and 20% mature segmented PMN, yielding a total of ±60% developing PMN.The length of time that PMN spend in each stage and in various compartments wasobtained mainly by radioisotope-labelling techniques (Athens et al., 1959; Cartwrightet aL, 1964). When the radioisotope diisopropyl fluorophosphate [32PJDF becameavailable, it provided a tool to estimate both the rate of PMN generation by the bonemarrow and their lifespan in the circulation (Athens et al., 1961; Athens et at., 1959;Cartwright et al., 1964; McAfee and Thakur, 1976). This isotope given systemically,labels all the granulocyte series of cells and labelling cells ex vivo labels only themature granulocytes. This difference has been used to estimate the rate of generationof PMN in the bone marrow and their lifespan in the circulation. By using thisisotope Cartwright et at (1964) show that the circulation half-life of PMN is ±7 hourswith a myelocyte generation time of ±3 days. Utilizing tritiated thymidine (H3TDR)to label cells engaged in DNA synthesis in vivo, it was shown that one coulddetermine the time from the last division in the myelocyte stage in the bone marrow5Figure 1: The lifespan and stages of maturation of polymorphonuclear leukocytes.The times for the various compartments were obtained by isotope-labelling technique.The ordinate shows the flux of PMN through each compartment and the abscissashows the time spend in each compartment. Note the stepwise increase in cellnumbers in the compartments where cell division took place. No mitoses occur afterthe myelocyte stage. The size of the tissue pooi of PMN is unknown.6to the appearance of labelled PMN in the peripheral blood (Bryant and Kelly, 1958;Fliender et al., 1964; Maloney, and Patt, 1968). Some similarities exist in the patternof release of PMN for different species; however, there are temporal differences. Theemergence time for segmented PMN into the peripheral blood ranges from 96-144hours in humans (Fliender et aL, 1964), 102±13.8 hours in dogs (Maloney and Patt,1968) and as short as 24 hours in mice (Bryant and Kelly, 1958). These emergencetimes (myelocyte-to-blood transit time) may vary considerably between patients andcould be explained by either a longer maturation time from the myelocyte to thesegmented PMN or a delay in the release of mature cells from the bone marrow. Thework of Cronkite et al (1960) and Maloney and Weber (1963) suggest that there is arandom release of cells into the blood from the marrow segmented pool of PMN incontrast to first-in first-out kinetics. Shorter emergence times in patients withinfections suggest that segmented PMN in these conditions are younger or moreimmature, compatible with the “shift to the left” or cytoplasmic “toxic granulation”seen on differential counts (Fliender et al., 1964). Whether these younger cellsreleased into the circulation have different kinetic and functional properties is stillunknown. Polymorphonuclear leukocytes spend only a small proportion of theirlifespan in the vascular space and circulating PMN turn over ±2.5 times per day toreplace the circulating pool. To accomplish this the bone marrow produces ±850 x 106cells/kg/day in humans. These cells come from the postmitotic granulocyte pool in thebone marrow (5.6 x iO cells/kg) which is the largest pool of PMN and is ± 10 timesthe size of the intravascular pool (Finch et al., 1977). Inflammation and stressincrease the rate of PMN production from the precursors, shorten the maturationtime, decrease the time mature PMN reside in the marrow, and stimulate mature7and immature PMN to enter the circulation (Cronkite, 1988). The myelocyte-to-bloodtransit time may be as short as 48 hours during infection (Fliedner et aL, 1964b). Theturnover of PMN may increase from 100 billion per day at baseline to over a trillionper day during a serious infection (Walker, and Willemze, 1980; Malech, 1988).b) Distribution of PMN in the vascular spaceThe fact that PMN can leave the circulation yet remain in the vascular space so thatthey can be rapidly mobilized into the circulation in times of stress was recognizedmore than a century ago by Virchow (1856), Cohnheim (1867) and Limbeck (Limbeck,1889). These early observations set the stage for a systematic investigation of thedistribution of leukocytes in the vascular system. Cohnheim established thatleukocytes can assume a marginal position in the venules, which was attributed tothe low blood velocity in the peripheral compared to the central vessels. Therelationship between blood velocity and margination provides the mechanism forleukocytes accumulate in some vascular regions and to be dislodged from others. Theimportance of the lung as the major site of PMN margination was first recognized byBruce and Andrewes (Andrewes, 19 lOb; Andrewes, 1910a; Bruce, 1894). Theydemonstrated that the redistribution of PMN from the circulation to the marginatedpool in the lung with the intravenous injection of peptone (Bruce, 1894), tubercie andtyphoid bacilli as well as Escherichia coli (Andrewes, 191Gb; Andrewes, 1910a).Goldscheider and Jacobs (1894) reached similar conclusions at about the same timeconcerning the importance of the lung as a site where PMN could withdraw to amarginal position. In the 1950’s, successful catheterization of the great vessels helpedto establish that PMN present in the vascular system could marginate along the8vessel walls in the lung with various stimuli that increase or decrease the numberof cells marginating within the pulmonary vasculature. Using these techniques,several investigators established that sequestration of leukocytes is a normalphysiological feature in the pulmonary circulation. Respiratory movements, such asthe Valsalva and Mueller manoeuvres either increase or decrease leukocytemargination respectively (Bierman et al., 1952a). Exercise and epinephrine infusiondemarginate leukocytes from the lung (Ahlborg and Ahlborg, 1970; Bierman et al.,1952b) which suggests that changes in pulmonary blood flow are an important factorinfluencing pulmonary sequestration of leukocytes. The concept that catecholaminesmay influence margination by decreasing the adhesive forces between PMN and theendothelial cells was suggested by Ahlborg et a! (1970) in a study showing that theleukocytosis of exercise can be blocked by the B-blocker, propanolol. However, thisfinding was subsequently challenged by Foster et a! (1986) who showed no effect ofB-blockers and emphasized the importance of bloodflow on PMN margination in thelung. A study by Thommasen et a! (1984) of the pulmonary arterio-venous differenceof PMN in dogs showed that PMN retention is flow-dependent, which supports theconcept of a close relationship between blood flow and PMN margination in thepulmonary circulation.The lung has unique anatomical and physiological features that are important indetermining whether PMN will move through the microvasculature or be retarded.In the larger conducting vessels, the PMN travel in the centre of the flowing streamand have few opportunities to interact with the endothelial surface. However, in thealveolar wall the PMN make close contact with the endothelial cells because the9mean diameter of the PMN (mean diameter 7-8 microns) is larger than manycapillary segments (mean diameter 5-6 microns) (Schmid-Schonbein et aL, 1980;Weibel, 1963). The PMN must pass through an average of 100 capillary segmentsfrom the arteriole to the venule and have been shown to deform as they pass throughthe capillary vessels. This results in intimate contact between the PMN and theendothelial membrane during their journey through the lung and in the PMN havinga much longer transit time than the red blood cells. This anatomical discrepancybetween PMN and capillary diameter was illustrated by Martin et al (1982) whofound that in dogs, 80 % of the PMN are removed from the circulation by thepulmonary capillaries in the first pass through the lung. Similar findings werereported by Muir et al (1984). They injected labelled PMN and erythrocytes (RBC) inupright awake humans, and found that ±20% of the labelled PMN failed to negotiatethe pulmonary vascular bed in the same amount of time as the erythrocytes.The transit time for PMN is longer than RBC and longer in the upper regions of thelung (Hogg et aL, 1985; Wagner et al., 1982). If the capillary pathways are the same(Staub and Schultz, 1968), the longer transit time in the flow independent lung areasmeans that the velocity of blood is slower in the upper regions of the lung, supportingthe concept that forces tending to propel the PMN and those retarding them can beshifted by changing blood velocity. This results in greater retention and a slowerturnover of PMN in the upper regions of the lung. This retention may cause a longerexposure of PMN to airborne activating stimuli, such as cigarette smoke, resultingin the typical distribution of smoke related lung emphysema (Leopold and Gough,1957).10Weibel et al (1963) has estimated that there are ±296 x 106 alveoli and 277 x iOcapillary segments in the human lung; therefore there are ±1000 capillary segmentsper alveolus. This capillary network allows large numbers of PMN to marginatewithout interfering with the majority of available pathways and with little effect onthe pressure drop across the pulmonary microvascular bed. The pulmonary capillarybed can accommodate large numbers of cells and calculations range from twice to 6times the circulating PMN pool (Doerschuk et al., 1987a; Staub et al., 1982).When radioisotopes became available with modern imaging techniques, they wereused extensively to advance the understanding of both the production and theintravascular behaviour of PMN. Most of the studies using isotopes have confirmedthe findings of many older studies concerning the distribution of PMN in thecirculation inthat injecting labelled PMN into the venous or arterial site of thecirculation results in delay in the lung with gradual accumulation in the liver andspleen over time (Martin et al., 1982; Muir et al., 1984; Saverymuttu et al., 1983).The influence of the ex vivo labelling techniques on the behaviour of the reinjectedPMN was recognized as an important factor in the discrepancies between differentstudies (Haslett et aL, 1985; Saverymuttu et al., 1983).c) Distribution of PMN in the tissueThe final tissue destination of most PMN in the normal host has not been resolved.Some PMN may migrate to areas of low-grade inflammation, such as the gingivalcrevasses around the teeth, upper respiratory tract and bladder, where they areimportant in host defense mechanisms (Van Dyk et aL, 1985). Earlier studies suggestthat the gastrointestinal tract is the most important site for PMN egress in normal11individuals (Murphy, 1976), but this finding has been questioned as few PMN areseen in this tissue in the absence of infection or other inflammatory processes(Jamuar and Cronkite, 1980). The reticuloendothelial tissue in the spleen and theliver may play a key role in the removal of senescent PMN from the circulation(Doerschuk et aL, 1987a; Doerschuk and Allard, 1989), however splenectomizedpeople do not undergo major change in the number of circulating PMN. Severalstudies have proposed that permanent removal of PMN from the intravascular spaceinto the tissues is a random process (Carper, 1966; Fliender et al., 1964; Raab et al.,1964), and that PMN never return to the circulation again. Like the site of removal,the signal for permanent removal of PMN from the circulation is unknown. The sizeof the tissue pool of PMN depends on the time they survive in this pooi before theyare processed. The fate of PMN after they have migrated into the tissues is stillunclear but they are reported to be capable of surviving for several days (Bainton,1980).122) BONE MARROW RELEASE OF PMN:The continuous supply of PMN to the circulating blood depends on their rate ofproduction (hematopoiesis) and their subsequent release into the marrow sinuses(hematoegression). Under normal circumstances only differentiated postmitoticgranulocytes cross the sinus wall to gain access to the circulation. The process bywhich cells gain access to the circulation is selective in several ways; for exampleonly mature cells that are at the end of the production line and prepared for theirfunctional role, egress from the bone marrow. Moreover, an increased appearance ofimmature white blood cells in situations such as infections (Boggs, 1967; Cronkite,1979), the administration of growth factors (Platzer, 1989) or corticosteroids (Cronkite,1979; Spertini et al., 1991b) implies unique or separate mechanisms for neutrophilproduction and release (Broxmeyer et al., 1974). The mechanisms that control therelease ofPMN from the bone marrow are poorly understood. Several factors probablyregulate the retention of immature cells in the bone marrow and the selective egressof mature leukocytes into the vascular space and out of the circulation into thetissues. Those that have been suggested include: the anatomical localization of thedeveloping hematopoietic cells adjacent to the vascular space (Lichtman and Erslev,1988); the development of transient holes or pores in the venous sinuses to allowegress of marrow cells (Weis, 1983); and developmental changes in the nucleus andcytoplasm of maturing PMN such as increased motility (Lichtman and Weed, 1972a),deformabiity (Lichtman, 1970), chemotactic ability (Giordano and Lichtman, 1973),that facilitate translocation of terminally differentiated PMN from the hematopoieticcompartment into the efferent vascular space. Releasing factors such as13glucocorticoid and androgenic steroids (Deinard and Page, 1974), endotoxins(Kampschmith, 1984), components of the complement system (Ghebrehiwet, andMuller-Eberhard, 1979), GM-CSF and G-CSF (Platzer, 1989) have also beenimplicated in the initiation of PMN release from the bone marrow but theirmechanism of action is still largely unclear.a) The structural and functional relationship of the bone marrow-bloodbarrierThe major supply of blood to the bone marrow is derived from the nutrient artery andpenetrating periosteal arteries. The ascending and descending central artery in thebone gives off radial arteries, becomes fine arterioles and ends in capillaries thatopen into the bone marrow venous sinusoids. These sinusoids begin at the endostealsurface and are fed primarily from intracortical capillaries. These vessels coursethrough the medullary cavity of the bone, anastomose, and eventually drain into alarge central venous sinus. These branching vascular sinuses (1O-3Om in diameter)may coalesce and form larger collecting sinuses (50-75jAm) before entering into thecentral sinus. The sinusoids are surrounded by fibroblastic stroma (Lewis, 1982) withthe hematopoietic tissue located in cordlike anastomosing bands between thesevenous sinuses. The hematopoietic activity of the marrow takes place in the centreof these cords and the cells that are generated begin to move towards the sinusoidsafter they have differentiated and enter the postmitotic pool (Weiss and Chen, 1975).Cells egress from the hematopoietic compartment of the bone marrow into thegeneral circulation by crossing the sinus wall.The sinus wall is a trilaminar structure (figure 2) consisting of a continuous layer of14endothellal cells, a discontinuous basement membrane and abluminal coveringadventitial reticulum cells (Weiss, 1970). The endothelial cells have several types offenestrae that are spanned by a diaphragm that is thought to open transiently toallow cells to move into the sinusoids (Muto, 1976). This suggests that the egressingcells pass through the endothelial cell and supports the observations of Tavassoli andShaldai (1979) who noted the absence of tight junctions in bone marrow endothelialsinuses. The fenestrae are thought to be dynamic structures that can be modulatedby regulating factors such as phorbol esters (Lombardi et al., 1987), and ACTH(Apkarian and Curtis, 1986) and may be controlled by the endothelial cytoskeleton(Steffan et aL, 1987). The basement membrane containing mainly laminin and typeIV collagen is discontinuous and completely lacking at the site of endothelialfenestrations (Campbell, 1972; Muto, 1976; Weiss, 1970). The adventitial cells thatcover the abluminal surface of the endothelial cells are fibroblastic in nature andproduce extracelluar matrix (ECM), collagen type I and Ill, reticular fibres andgranules containing material resembling the ECM (Bentley and Foidart, 1980; Weiss,1976). The adventitial cell is also rich in microtubules (Weiss, 1976) that allow themto retract their extensions and change the percentage covering of the endothelial cells(called the adventitial cell cover rate or ACCR). The normal ACCR is 60 %-65 %(Weiss, 1970) and altered in situations of increased marrow cell egress such as afterendotoxin treatment (Weiss, 1970), post-phlebotomy (Tavassoli, 1977) or in myeloidleukemias (Leonardi, and Manthos, 1989; Nagaoka et al., 1986). Egressing blood cellscannot pass through the adventitial cells but have to move between or around them(pericelluar passage). Although the functional significance of the adventitial cells inregulating cell egress is still unclear it has been suggested that these cells regulate“5Bone Marrow Blood-barrierAdventitial Cell Mature PMNBasaJL0Figure 2: Discontinuous bone marrow-blood barrier. Mature PMN egressing from thehematopoietic compartment into the bone marrow venous sinusoids have to passthrough three layers (adventitial cell layer, basal lamina, and endothelial cell).IrFenestraEndotheijal Cell16the selectivity of the bone marrow-blood barrier (Petrides and Dittmann, 1990).In addition to the stromal and accessory cells, the microenvironment of the bonemarrow contains soluble molecules and extracelluar matrix. The ECM is composedof collagen type I and ifi (made by fibroblasts), type IV (made by endothelial cells)(Reilly et al., 1985), glycoaminoglycans, proteoglycans and glycoproteins such asfibronectin (Bentley et aL, 1981). The interactions between the ECM and thehematopoietic cells are crucial for the proliferation and differentiation of thehematopoietic cells (Owen, 1988). Currently little is known about the interactions ofegressing neutrophils and the ECM and stromal cells. Some have speculated that theECM serves as an anchor reducing the discharge of immature cells from the bonemarrow (Lichtman, 1981), and that it provides a barrier with a high degree ofplasticity (i.e. that it behaves as a reversible deformable meshwork that can bemechanically displaced by the cell having the capacity to egress). Membraneassociated proteins on white cells and their ligands on the ECM could be responsiblefor these interactions (see below). Anatomical pathways for cell movement across thebone marrow-blood barrier may be responsible for a constant “leak” of hematopoieticelements into the sinusoids under stable condition. However there is a growing bodyof evidence favoring the hypothesis that the egress of cells from the bone marrow isan active process rather than a passive leakage of cells through endothelial pores(Petrides and Dittmann, 1990). This active process probably involves changes in thehematopoietic cells and their microenvironment that allow immature cells to attachto the ECM during differentiation and mature cells to detach from the mathx andcrawl into the sinusoids.17b) Developmental changes and leukocyte egress from the bone marrowThe production of granulocytes involves a variety of different hematopoietic growthfactors. Colony-forming units for granulocytes and monocytes (CFU-GM) proliferateand differentiate under the influence of specific hematopoietic growth factors, notablygranulocyte-monocyte colony stimulating factor (GM-CSF) and IL-3, which areproduced locally in the bone marrow microenvironment by endothelial cells,fibroblasts and augmented by circulating T-lymphocytes and monocytes. Over thecourse of 5 to 7 days they become committed to either neutrophil or monocytedifferentiation. At this point in their differentiation these cells are largely regulatedby the local concentration of growth factors specific for their individual lineages,granulocyte colony stimulating factor (G-CSF) for neutrophils and monocyte colonystimulating factor (M-CSF) for monocytes. Hematopoietic stroma (stromal cells andconnective tissue) play a pivotal role in this local production of growth anddifferentiation factors for granulocytes. These cells secrete interleukins, colonystimulating factors and monokines that serve as growth and signaling factors eitherby acting directly or indirectly on hematopoietic cells. IL-i, for example, has nocolony stimulating activity itself but, when administrated in vivo, universally inducesa neutrophilic leukocytosis, which results from the induction of G-CSF and GM-CSFexpression by other accessory or auxiliary cells such as fibroblasts (Lee et al, 1987)and endothelial cells (Bagby et a!, 1986). IL-6, secreted by bone marrow macrophages,fibroblasts and endothelial cells, has no direct effect on proliferating humanhematopoietic cells but functions synergistically with many direct actinghematopoietic growth factors in promoting hematopoiesis (Ogawa et al, Blood Cells1988). This clearly illustrates the the dynamic interactive relationship between the18bone marrow stromal cells, connective tissue and hematopoietic cells. The final phaseof myelopoiesis, from the myeloblast through promyelocyte, myelocyte,metamyelocyte, band forms and segmented neutrophils accompany another 4 to 8divisions and occur over 7 to 10 days. The growth factors and interleukins thatinduce proliferation and differentiation of granulocyte precursor cells often enhancethe functional activity of their terminally differentiated progeny (Clark and Kamen,1987, Gasson et al, 1984, Mayer et al, 1987), that may include promoting egress fromthe bone marrow.These hematopoietic growth factors induce developmental changes in the nucleus andthe cytoplasm of maturing cells which play a pivotal role in allowing thetranslocation of terminally differentiated cells from the hematopoietic compartmentto the efferent vascular space. Marked increase in mobility (Giordano and Lichtman,1973; Lichtman and Weed, 1972b) deformability (Lichtman and Weed, 1972a) andchemotactic ability (Giordano and Lichtman, 1973) occur during the maturation ofthe granulocytes. Active motility is not a feature of immature hematopoietic cells andthe ability of mature granulocytes to probe by pseudopods probably contributes totheir ability to search out the sinus wall and penetrate it (Giordano and Lichtman,1973). The ability of cells to deform relates to their ability to move. Deformabiity ofhematopoietic cells of different stages of maturation has been examined by theirability to be filtered through micropore membranes (Lichtman and Kearney, 1976)and to be aspirated into glass microcapillary tubes (Lichtman and Weed, 1972a;Lichtman and Weed, 1972b). Mature granulocytes can migrate through filters with1m pores. The biophysical character of maturing cells is important in marrow egressbecause most cells undergo marked deformation during migration through pores in19the sinus wall. Like motility, chemotaxis of granulocytes is a feature of maturerather than immature cells (Giordano and Lichtman, 1973). Chemotaxis is importantbecause it may underlie the ability of mature marrow cells to accelerate their releasein response to periods of increased demands. All these abilities combined maycontribute to allowing the PMN to cross the bone marrow-blood barrier. Themigrating cells are thought to make the pores that develop in the endothelial cellcytoplasm (Giordano and Lichtman, 1973). These holes frequently occur adjacent tojunctions of endothelial cells with the majority of cellular traffic migrating throughendothelial cells.c) Factors that cause release of PMN from the bone marrowCell releasing factors could affect the rate of egress of PMN from the bone marrowin several ways. Firstly, they may act as specific chemoattractants that acceleratemovement of differentiated cells into the marrow sinusoids. Secondly, they could acton the sinus wall and stromal structures to reduce the impediment to egress byreducing the adventitial cell cover rate (Weiss, 1970) which demonstrated withendotoxin induced bone marrow release of leukocytes (Weiss, 1970). Changes in theadhesion properties of leukocytes to bone marrow structures by factors that inducebone marrow release of leukocytes may provide new insights into this complexproblem.Well characterized releasing factors for granulocytes included: components of thecomplement-system (Ghebrehiwet and Muller-Eberhard, 1979); glucocorticoid andandrogenic steroids (Deinard and Page, 1974); endotoxins (Kampschmith, 1984; Voset al., 1972); GM-CSF; and G-CSF (Platzer, 1989). Complement fragments that have20been shown to induce bone marrow release of leukocytes are cleavage products of C3such as C3a and C3e (Ghebrehiwet and Muller-Eberhard, 1979). Purified C3e wasfound to mobilize leukocytes from bone marrow upon perfusion of isolated rat femurs.The observation that an individual with homozygous C3-deficiency did not respondwith a leukocytosis during many episodes of infection (Alper et aL, 1972) suggeststhat C3 is critical in the complement-dependent release of leukocytes from the bonemarrow. Complement fragments such as C5a, as well as lipopolysaccharides (LPS)and GM-CSF, have also been shown to activate PMN and induce shedding of the celladhesion molecule, L-selectin, from the PMN surface (Griffin et aL, 1990). Theassociation between a possible de-adhesive events such as the shedding of L-selectinand the bone marrow release of PMN raises the possibility that the shedding of Lselectin contributes to the relocation of more immature granulocytes from the bonemarrow to the circulating blood.d) Adhesion events and egress of PMN from the bone marrowThe control of cell-cell and cell-matrix interactions via the cell adhesion proteins onthe surface of hematopoietic cells has emerged as an important pathway for theregulation of cell movement and migration (Campbell and Wicha, 1988; Hynes, 1992;Springer, 1990b). These adhesion interactions are thought to play a crucial role inthe regulation of hematopoiesis and the direction and control of leukocyte traffic,egress from the bone marrow into the circulation and migration into tissues(Campbell and Wicha, 1988; Williams et al., 1991; Simmons et al., 1992; Miyake etal., 1990a). The mechanisms involved in the coordinated egress of specific cell typesfrom the bone marrow require that the barrier be both selective and responsive to21stressful stimuli. Both cell-matrix and cell-cell interactions, as well as solublemediators, may be involved in the control of this marrow-blood barrier selectivity.d.1 Cell-matrix interactions:The spatial proximity of hematopoietic cells and local sources of growth factors maybe largely regulated at the level of cell contacts. The interaction of granulocytes withstromal components may also be important for retaining precursors in the bonemarrow. Studies of peripheral leukocytes have revealed a number of molecules thatmediate cell-cell and cell-mathx interactions. Recent reports indicate that some of thesame cell adhesion molecules mediate the binding of myeloid progenitor cells to bonemarrow stromal elements (Miyake et al., 1990a; Simmons et al., 1992). There isample evidence to show that bone marrow cell-extracellular matrix interaction occursand that this interaction is important for hematopoiesis (Miyake et al., 1990b).Monoclonal antibodies to CD44 block lympho-hemopoiesis in long-term bone marrowculture (Campbell et al., 198Th; Wolf, 1979; Dexter, 1982). Whether changes in theseadhesive interactions are involved in the egress of cells from the bone marrow is lessclear. Fibronectin is a major component of the ECM that is present on the stromalcell membranes of the bone marrow (Torok-Storb, 1988; Tsai et al., 1987; Weiss andReddi, 1981). Erythroid progenitors and precursors remain bound to the cell-bindingdomain, the universal attachment peptide arginine-glycine-aspartic acid (RGD), onfibronectin throughout differentiation (Tsai et al., 1987). Loss of receptors from theerythroid progenitors for the RGD domain on fibronectin is believed to initiate therelease of reticulocytes from the marrow (Pate! and Lodish, 1984; Vuillet-Gaugler etal., 1990). Membrane associated chondroitin sulphate (CS) on hematopoieticprogenitors can also mediate the binding of these cells to fibronectin on stromal cells22via the RGD motif (Ruoslahti, 1989; Minguell et aL, 1992). Although most studieshave focused on the role of proteoglycans or glycoaminoglygans in themicroenvironment to modulate proliferation of hematopoietic cells, at least one studysuggests a role for these ECM components in the adhesion of myeloid precursors (DelRosso et aL, 1981). This study found that mature granulocytes were unable to adhereto preformed marrow stromal cells, but that treatment of these granulocytes withhyaluronidase restored 60 % adhesion capability to marrow stromal cells in vitro.Immature granulocytes have little glycoaminoglycans on their surface and membranebound glycoaminoglycans on mature cells were capable of interrupting adhesiveinteractions between the mature granulocytes and the marrow stroma. However, thespecificity of this interaction was not strictly evaluated.It has been suggested that the interaction of the hematopoietic isoform of CD44(CD44h) with the glycoaminoglycan hyaluronate to anchors immature hematopoieticcells to bone marrow ECM (Carter, 1982; Miyake et aL, 1990a; Stamenkovic et aL,1991). Expression of CD44 correlates with maturation status among bone marrowprecursors and primitive myeloid progenitors cells are CD44M while expressiondecreases on more committed granulocytes (Kansas et a!., 1990; Lewinsohn et al.,1990). Reduced interaction between cellular CD44 and ECM hyaluronate duringgranulocyte differentiation and maturation may allow these cells to translocate tosinus walls in the bone marrow. Mature granulocytes, however, upregulate CD44 ontheir surface before egressing into the circulation. The possible role of thesemature granulocytes in bone marrow egress is still unknown.Campbell et a! (Campbell et al., 198Th; Campbell et al., 1987a) isolated an ECM23protein of relative molecular weight 6Okd that mainly attached to cells from thegranulocyte lineage and called it hemonectin. This molecule was found only in thebone marrow, suggesting that it is both an organ-specific and a lineage-specific ECMcomponent and providing a possible molecular mechanism for granulocyticpredominance of bone marrow compared with other hematopoietic tissue such as thespleen. The adhesion of immature bone marrow cells but not circulating granulocyticcells to hemonectin suggests that developmentally regulated adhesion to hemonectinmay be involved in the mechanism of release of granulocytes from marrow (Campbellet aL, 198Th). The factors regulating the interaction between hemonectin andimmature granulocytes in the bone marrow or the counter receptor on these cells forhemonectin are currently being investigated.d.2) Cell-cell interactions:Lectins mediated binding in the bone marrow:Lectins on cell surfaces mediate cell-cell interactions by combining withcomplementary carbohydrates on apposing cells and play a key role in the control ofvarious normal and pathological processes in living organisms (Brandley andSchnaar, 1986; Lis and Sharon, 1986; Sharon and Lis, 1989a; Liener et aL, 1986) suchas fertilization, embryogenesis, immune defense, cell migration and microbialinfection. In the 1970’s it became well-established that almost all cells carrycarbohydrates on their surfaces in the form of glycoproteins or glycolipids (Cook,1986). The lectins are glycoproteins that bind carbohydrates specifically, noncovalently and with speed and reversibility (Cook, 1986; Lis and Sharon, 1986;Sharon and Lis, 1989a; Liener et al., 1986). Typically, the lectins and thecomplementary carbohydrates are located on the surfaces of apposing cells, which24may be of the same type or of different types. Furthermore, cell surface lectins maycombine with carbohydrates of the ECM that promote cell-matrix adhesion. Thephenomenon of clinical bone marrow transplantation, in which stem cells lodge andproliferate preferentially within the bone marrow, results from specific adhesion ofthese stem cells to macromolecular components of ECM or on the stromal cells uniqueto bone marrow. This binding of progenitor cells to stromal cells and ECM involvesan interaction between a membrane lectin on the one side and a membrane or ECMglycoconjugate on the other (Tavassoli and Hardy, 1990). Matsuoka et al (1989)purified a lectin homing protein (MW 110,000) from cloned progenitors withcarbohydrate-binding qualities, suggesting that a lectin on progenitors is involved inthis interaction. This supports a wealth of recent information on membrane lectinswith biological recognition functions (Hoppe and Lee, 1982; Lehrman and Hill, 1986;Prieels et aL, 1978; Thornburg et aL, 1980). These membrane lectins are calciumdependent in their binding to carbohydrates and are known as C-lectins (Drickamer,1988; Drickamer and McCreary, 1987). Similarities exist between this homing proteinin hematopoietic progenitors and the homing receptors on lymphocytes and PMNbelonging to the selectins group of adhesion molecules (Stoolman et al., 1984;Yednock and Rosen, 1989; Butcher, 1990). These molecules have been shown to beinvolved on lymphocyte- and neutrophil-endothelial cell interactions.Granulocytopoiesis involves differentiation of bone marrow stem cells into thegranulocyte cell lineage and maturation of the granulocyte-committed cells, whichundergo a limited number of cell divisions and finally enter the circulation assegmented granulocytes. Each step in this process is likely influenced by interactions25between granulocyte precursors and the bone marrow microenvironment.Furthermore, during bone marrow release of PMN, cells have to cross an endothelialbarrier when moving from the hematopoietic tissue into the bone marrow venoussinusoids. The cell-cell and cell-matrix interactions that are involved and that controlthis process are largely unknown. Cell-cell interactions via the cell adhesionmolecules are important for leukocytes to cross the endothelial barrier to sites ofinflammation in the systemic and pulmonary vascular beds (Butcher, 1991; Butcher,1990; Lawrence and Springer, 1991; Yong and Khwaya, 1990). The initial cell-cellcontact between PMN and endothelium is mediated by the selectin family of adhesionreceptors. One of these selectins, L-selectin, is constitutively expressed by nearly allintravascular granulocytes and interacts with the other inducible selectins, P- andE-selectin, as well as inducible carbohydrate moieties on the endothelium to initiateleukocyte-endothelial adhesion. L-selectin is shed from the surface of leukocytesduring recruitment to foci of inflammation (Humbria et al., 1994; Julita et al., 1989;Kishimoto et al., 1989; Porteu and Nathan, 1990; Tedder, 1991; Spertini et al.,1991b). For example, in patients with rheumatoid arthritis, L-selectin expression onPMN in synovial fluid is much lower than expression on circulating PMN thatimplies shedding of L-selectin when PMN emigrate from the vascular space (Humbriaet al., 1994). It has been proposed that L-selectin shedding during PIvIN activationmay be a de-adhesion event and a signal for migration of the PMN through theendothelial cell barrier and into the underlying tissue (Tedder, 1991; Kishimoto etal., 1989; Jutila et a!., 1989; Smith et al., 1991).In the bone marrow, L-selectin is expressed on nearly all postmitotic myeloid cells(Griffin et a!., 1990), and Lund-Johansen et a! (Lund-Johansen and Terstappen, 1993)26have shown that L-selectin expression increases with granulocyte maturation in thebone marrow. The fact that the crossing of an endothelial barrier is associated withL-selectin shedding led us to consider the possibility that a similar shedding of Lselectin occurs when PMN cross the bone marrow-blood barrier.273) SELECT1NSLectin or carbohydrate recognition domains of proteins have been shown to beinvolved in a number of biological events (Sharon and Lis, 1989b), but therelationship between carbohydrate-mediated cell adhesion and leukocyteinflammation was not always clear. The discovery of the selectins, a family of threecell surface glycoproteins containing lectin domains that mediate regionalinflammation responses by recognition ofcell-specific carbohydrates, has unified thesedivergent fields and has given rise to some observations that may ultimately proveto be clinically significant. The selectins are a group cell adhesion molecules includedamong an increasing number of leukocyte adhesion molecules that act in acoordinated manner to orchestrate the migration of various classes of leukocytes,during lymphocyte recirculation, inflammation, metastasis and other types ofleukocyte traffic (Hemler, 1988; Springer, 1990b). Recent work has demonstrated theexistence of this novel family of adhesion molecules that appear to utilize proteincarbohydrate interactions for specific cell-cell binding between leukocytes andendothelial cells. Molecular cloning revealed the unifying aspect of this new familyof three adhesion molecules to be their common structure, consisting of a calciumdependent lectin- or carbohydrate binding domain, an epidermal growth factor typemotif (EGF), variable numbers of complement regulatory moieties (CR), atransmembrane region and a cytoplasmic tail (Bevilacqua et al., 1989; Geng et al.,1990; Stoolman, 1989; ) (figure 3). The most notable is the C-type lectin motif foundat the N-terminus of the protein which has a 60 % -70 % amino acid sequence homologybetween the three different selectins and functions as the principal component in celladhesion events (Bevilacqua et al., 1989; Johnston et al., 1989; Lasky et al., 1989;28Tedder et al., 1989; Siegelman, and Weissman, l989b; ). While the lectin domainseems to be the major determinant of cell adhesion, its structure and function appearto be conformationally dependent upon the EGF and the CR domains (Bowen et aL,1990; Watson et al., 1991). L-selectin (CD62L) is expressed on nearly all leukocyteswhile E-selectin (endothelial-leukocyte adhesion molecule or ELAM-1, CD62E) andP-selectin (CD62, PADGEM, GMP-140, CD62P) are inducible molecules on theendothelial cell surface. Table I summarizes the properties of the selectins.L-selectinL-selectin is the smallest cell adhesion molecule of the selectin family and is thehuman counterpart of the murine lymphnode homing receptor identified by the MEL-14 antibody (Gallatin et aL, 1983a). L-selectin was first identified structurally bycDNA cloning (Bowen et al., 1989; Siegelman et al., 1989a; Tedder et al., 1989), islocated on chromosome 1 at band q23-25. The three selectin genes, are within 200kilobases of one another in the human genome, suggesting that they derived from anevolutionary related gene (Collins et al., 1991; Ord et al., 1990; Watson et al., 1990).It is noteworthy that the selectins genes map very close to the locus on the genes thatcode for complement activation proteins (such as complement receptor 1 and factorB) with the same complement repeat motif that is found in the selectins. It is likelythat this family of adhesion molecules evolved from a common progenitor genethrough gene duplication and exon amplification.1) Structure of the L-selectin protein: (figure 3)The L-selectin on lymphocytes is smaller (Mr of ±74,000) than the molecule on PMN2qSelectins: Nomenclature, expression and functionName Cell type Expression Proposed functionL-selectin Lymphocytes Shed after cell LymphocyteLECAM-1 Monocytes activation(min). recirculation-PLN.LAM-i Neutrophils Conformational Leukocyte rolling.TQ-1 Eosinophils change(?). NeutrophilLeu-8 inflammation.CD62LMEL-i4P-selectin Platelets Increases upon Neutrophil/monocytePADGEM (cr-granules) thrombin, rolling.GMP- 140 Endothelium histamine, LeukocyteCD62P (W-P bodies) Substance P inflammation.activation(min).E-selectin Endothelium Increases upon Neutrophil/monocyteELAM-1 transcriptionally IL-i, TNF, LPS rolling.CD62E activated activation(hours). Leukocyteinflammation.PLN, peripheral lymph nodes; ELAM-1, endothelial-leukocyte adhesion molecule 1;GMP-140, granule membrane protein 140; PADGEM, platelet activation-dependentgranule external membrane protein. TNF, Tumor necrosis factor; LPS,lipopolysaccharide; W-P, Weibel-Palade.Table I30(Mr of 90,000 to 100,000) (Griffin et al., 1990) and both molecules are highlyglycosylated as L-selectin cDNA encodes only a ±37kd, protein suggesting post-translational processing of the molecule. The molecule has a C-type lectin domain,a short epidermal growth factor-like domain, two complement moieties homologousto those found in C3/C4 binding proteins, a membrane spanning domain and acytoplasmic tail (figure 3). The lectin-like domain of L-selectin is homologous withother carbohydrate binding proteins, including asialoglycoprotein receptors andmannose binding proteins. This domain contains essentially all the invariantresidues found in C-type amid lectin carbohydrate recognition domains (Drickainer,1988). Both murine and human homing receptors have been shown to required Cato function (Stoolman et al., 1987).The EGF domain contains the 6 Cys-residues conserved in all EGF domains and is39% identical to the amino-acid sequence of EGF. As EGF domains in severalproteins are involved in mediated protein-protein interactions, the L-selectin EGFdomain may serve a similar function and may not be merely a structural feature ofthe molecule (Kansas et at, 1991). The two complement motifs in L-selectin are theminimum protein binding unit in the C3/C4 binding proteins and the conservationof these moieties signals possible function significance in binding interactions.One striking feature between the lymphocyte homing molecule MEL-14 and Lselectin on other leukocytes is the homology in the transmembrane domains and thecytoplasmic tail, which is 88 % homologous. These regions in E- and P-selectin are nothomologous to L-selectin. L-selectin is shed from the cell surface following cellularL-selectjnN COO H31P-seled inNH2_f- Co OHE-selectjnNH2-{___- COOHFigure 3: Domain compositions of the three known selectins. The extracellularportion of each selectin contains an amino-terminal domain homologous to the C-typelectins, an adjacent epidermal growth factor domain followed by a variable numberof complement regulatory domains (circles) and transmembrane sequence (blackdiamond). A short cytoplasmic tail (open rectangle) is at the carboxyl terminus ofeach selectin.32activation, which makes it likely that this region is of critical importance in thisphenomenon and thus in the regulation of L-selectin expression. The cytoplasmic tailof L-selectin contains two potential sites for phosphorylation by protein kinase C.Phosphorylation of the tail has been postulated to be associated with the shedding ofthe molecule from the surface of leukocytes (Kansas et al., 1991). However, directbiochemical evidence of phosphorylation has been difficult because agents thatstimulate phosphorylation also enhance L-selectin shedding.2) Expression of L-selectin on leukocytes:The expression of L-selectin, which is limited to hematopoietic cells (Table I), is acomplex function of lineage, stage of differentiation, activation status, and anatomicallocation of the cells. L-selectin is expressed on the surface of most leukocytesincluding lymphocytes, neutrophils, monocytes, eosinophils, hematopoietic progenitorscells and immature thymocytes (Griffin et aL, 1990; Tedder et aL, 1990a). L-selectinis expressed late in B-cell development, is lost after activation, and is re-expressed onmemory B-cells. Similar patterns hold for T-cells, it is expressed on virgin T-cells, lostduring cell activation and re-expressed by most memory T-cells (Tedder et aL, 1985;Tedder et aL, 1990b; Kansas et al., 1985b; Kansas et al., 1985a). Early erythroidprogenitors cells (BFU-E) are L-selectin positive with a lack of expression on maturenucleated or non-nucleated erythrocytes.Among myeloid cells, L-selectin is expressed by nearly all circulating neutrophils,monocytes, and eosinophils (Griffin et aL, 1990; Tedder et aL, 1990a). The largestportion of L-selectin is found on the cell surface and does not localize in the33intracellular compartment. In contrast to lymphocytes, mature neutrophils andmonocytes, express low levels of mRNA for L-selectin, which suggests messageinstability or different post-translational protein processing in myelomonocytic cells(Tedder et al., 1989; Ord et al., 1990). L-selectin is expressed continuously throughoutmyeloid differentiation in the bone marrow (Griffin et al., 1990; Kansas et aL, 1990),including 80 % of CFU-GM that is the earliest progenitor cell committed to thegranulocyte/monocyte differentiation. The role of L-selectin in the localization ordevelopment of progenitors is currently unknown. As progenitors are known not totraffic into lymphnodes or inflammatory sites, it is possible that L-selectin maymediate leukocyte/extracellular matrix (ECM) or endothelial interactions within thebone marrow. Furthermore, the function of L-selectin on myeloid cells at the differentstages of differentiation and possible changes with maturation is still unclear.3) Regulation of L-selectin expression:An interesting and unique feature of the regulation of L-selectin expression is theloss of the molecule from the surface of leukocytes following cellular activation.Treatment of leukocytes with a variety of stimuli (Kishimoto et al., 1989), such asphorbol esters, complement fragments, tumor necrosis factor, leukotrine B4 and LPS,reduced the cell surface expression of L-selectin on T- and B-cells as well as ongranulocytes (Tedder et al., 1990a). A large fragment of L-selectin (Mr ±69,000) canbe immunoprecipitated from the supernatant, demonstrating that L-selectinexpression is down-regulated by shedding rather than by internalization, which isdistinct from that which down-regulates modulation of most other surface molecules.It has been proposed that the shedding of lymphocyte L-selectin might be necessaryto enable these leukocytes to transmigrate through the endothelium into sites of34inflammation, providing a rapid means for the adhesion and de-adhesion ofleukocytes to the endothelium (Kishimoto et al., 1989; Jutila et aL, 1989). Anotherpossible signal for receptor shedding is ligand binding which would provide a rapidmeans for leukocyte adhesion to the endothelium with subsequent de-adhesionfollowing receptor shedding. The temporal relationship between leukocyte-endothelialinteraction during inflammation and L-selectin shedding has not been established.Although the mechanism of shedding of L-selectin remains unknown, it has beenproposed that a membrane-bound protease with chymotrypsin activity cleaves thereceptor near the membrane, releasing a nearly intact extracellular domain withligand binding activity (Spertini et al., 1991b; Schleiffenbaum et al., 1992). This doesnot appear to result from the activation of induced soluble proteases released byleukocytes as the supernatant from the fluid of cells that have shed L-selectin doesnot contain soluble proteases that can alter the expression of L-selectin. Therefore,it is likely that enzymatic cleavage of L-selectin may result from the specificactivation of a membrane bound protease that may be rather ubiquitous since a broadarray of L-selectin negative cell types transfected with L-selectin cDNA are able toshed the receptor (Spertini et al., 1991b). The function of L-selectin may be furtherregulated by the presence of functional L-selectin in the extracellular environment(Schleiffenbaum et al., 1992). Soluble L-selectin in normal plasma caused a small butconsistent inhibition of lymphocyte attachment to high endothelial venules andhigher concentrations completely inhibited L-selectin-dependent leukocyte attachmentto endothelium. This soluble serum L-selectin may be a protective mechanism toreduce random leukocyte recruitment during generalized intravascular leukocyte35activation.A small amount of this soluble isoform of L-selectin (sL-selectin) can also beimmunoprecipitated from the supernatant fluid of lymphocytes cultured withoutstimulation, suggesting that L-selectin is constitutively shed at a slow rate withexpression kept constant by the re-synthesis of a new receptor (Spertini et al., 1991b).The demonstration of sL-selectin in circulating human plasma by Schleiffenbaum etal (1992) supports the idea that circulating leukocytes constitutively shed L-selectinintravascularly. The relatively high levels of soluble L-selectin in plasma and therelationship to the circulating leukocyte count suggest a possible role of sL-selectinregulating the circulating leukocyte count and leukocyte recruitment (Schleiffenbaumet al., 1992).4) Ligands for L-selectin:The ligand used by lymphocyte L-selectin (MEL-14) to home to peripheral lymphnodesvia high endothelial venules has recently been identified as carbohydratedeterminants Sgp5O and Sgp9O, coined GLYCAM-1 (Lasky et al., 1992b). Thiscarbohydrate ligand has been shown to be fucosylated, sulfated and sailylated, allcomponents essential for ligand binding activity to L-selectin (Ley et al., 1991a; Trueet al., 1990). The ligand(s) or counter receptor for L-selectin in postcapillary venulesof non-lymphoid tissue has yet not been identified. However, in vitro findings havesuggested that the L-selectin on PMN can interact with inducible endothelial celladhesion molecules by presenting sialylLewisx (sLex) and related carbohydrates asligands to the vascular selectins, P- and E-selectin (Picker et al., 1991). Mulligan et36al (1993) have demonstrated that synthetic sialyl-Lewis X protects against P-selectindependent lung injury, confirming the in vivo importance of this interaction.Moreover, the congenital absence of sLex ligand results in the clinical syndrome ofleukocyte adhesion deficiency and an inability of PMN to roll on activatedendothelium (Jutila et al., 1991; Segal et al., 1976). Furthermore, L-selectin, like theother selectins, has been shown in vitro to interact with sLex through its lectindomain (Foxall et al., 1992), or other still unidentified ligand(s) expressed on cytokinestimulated human umbilical vein endothelium (Spertini et aL, 1991a). NorgardSumnicht et al (Norgard-Sumnicht et aL, 1993) have identified in non-lymphoidendothelium a heparin-like ligand that binds a L-selectin chimera molecule; andrecently Baumhueter et al (1993) established that the protein core of Sgp9O isidentical to the sialomucin molecule CD34, suggesting that a glycoform of CD34 couldfunction as a ligand for L-selectin. This is relevant considering the broader tissuedistribution of CD34 with apparent constitutive expression on the endothelium in adiversity of non-lymphoid blood vessels (Fina et al., 1990). Differential vessel-specificglycosylation and translocation to the endothelial surface may be potential pathwaysof regulating L-selectin-CD34 interaction during the acute or chronic inflammatoryresponse. It remains to be seen whether CD34 is subject to biosynthetic control viacytokines, as has been reported for the ligand for L-selectin in cultured endothelialcells (Delia et at, 1993; Spertini et al., 1991a).5) Functions of L-selectin:As observed by Cohnheim (Cohnheim, 1889) more than a 100 years ago usingintravital microscopy, leukocytes begin to interact with the vessel wall by rolling37along the endothelium to adjacent tissue within minutes after injury. The rollingresponse is seen throughout the vertebratae, in both cold-blooded animals such asamphibians and in mammals (Cohnheim, 1889). The number of rolling leukocytesincreases dramatically during the course of an inflammatory reaction (Atherton andBorn, 1972) and is important in the accumulation and emigration of leukocytes inposteapillary venules (Fiebig et aL, 1991). The velocity at which cells tumble and rollin shear flow near the vessel wall is much faster than what is observed for rollingcells on inflamed endothelium, suggesting that an enhanced adhesive interactionoccurs between the leukocyte and the vessel endothelium (Atherton, and Born, 1973).The selectin family of cell adhesion molecules is specialized to mediate the rolling ofleukocytes and rolling observed in vivo during inflammation may involvecontributions from all three selectins. In PMN, L-selectin contributes to theinteraction of inactivated cells with cytokine-stimulated endothelial cells atphysiological shear stresses (Smith et aL, 1991). The other two selectins, in contrast,are cytokine-inducible cell adhesion molecules sequentially expressed on endothelialcells to support rolling of leukocytes, P-selectin being up regulated within minutesand E-selectin within hours (Abbassi et al., 1993; Springer, 1990a). In this cascadeof events PMN L-selectin plays a central role, functioning simultaneously as aconstitutively functional lectin recognizing molecules that bind to inducibleendothelial cell surface ligands, and as a structure presenting sLex to both P- and Eselectin whose expression is induced by inflammatory stimuli (Picker et al., 1991;Spertini et al., 1991a; Von Andrian et al., 1993). Evidence that this rollingphenomenon mediated by the selectins is essential for PMN emigration to areas ofinflammation comes from a limited number of animal studies (Ley et al., 1991b;38Mulligan et al., 1991; Mulligan et aL, 1994; Watson et aL, 1991). Monoclonalantibodies to L-selectin and a L-selectin-IgG chimera inhibit PMN rolling inmesenteric blood vessels and peritoneal inflammation (Watson et al., 1991; Ley et al.,1991b; Von Andrian et aL, 1991); monoclonal antobodies to E-selectin inhibit PMNmediated damage to the vascular endothelium during acute inflammation of the lung;and P-selectin chimera protect against complement mediated acute lung injury(Mulligan et aL, 1993; Mulligan et al., 1994). L-selectin also augmented intravascularaggregation of PMN in conjunction with theI2-integrins (Simon et aL, 1992). Thesefunctions of PMN L-selectin underline the importance of this molecule during theinitial phase of the inflammatory response.The work by Gowans et al (Gowans and Knight, 1964) suggested tissue-specificadhesive interactions of lymphocytes that home back to sites from which they wereinitially derived, which led to the identification of the lymphocyte L-selectinrecognized by the MEL-14 mAb that directed lymphocyte homing to post-capillaryhigh endothelial venules of peripheral lymphnodes (Gallatin et al., 1983b; Gallatinet aL, 1983a). In addition to directing lymphocyte recirculation through peripherallymphnodes, L-selectin on lymphocytes may also function in a similar fashion to Lselectin on PMN in recruiting lymphocytes to foci of inflammation (Lasky, 1992a;Michie et al., 1993).394)L-SELECTIN EXPRESSION DURING ACTIVE BONE MARROW RELEASEOF POLYMOPHONUCLEAR LEUKOCYTESIntroductionThe emigration of polymorphonuclear leukocytes (PMN) at sites of inflammationrequires intercellular adhesion. In the last 7 years an extraordinary increase ininformation concerning the mechanisms that mediate this leukocyte-endothelial cellinteraction during inflammation has become available (Butcher, 1991; McEver, 1992Springer, 1990b; Zimmerman et aL, 1992; Yong, and Khwaya, 1990). Duringinflammation leukocyte adherence is observed in postcapillary venules whereleukocytes roll on the luminal surface of endothelial cells before stopping. Theleukocytes then spread and adhere more tightly and finally emigrate into theperivascular tissue. Evidence indicates that the adhesion molecules supporting therolling phenomenon are distinct from the those required for stopping andtransmigration. This transient adhesion of leukocytes to regionally activatedendothelium is mediated by the selectins (Lasky, 1992a). The selectins are threerelated membrane proteins that interact with cell-surface carbohydrate ligands. Oneof these selectins, L-selectin, is expressed on most leukocytes, binds to inducibleligands on the endothellum that loosely adhere leukocytes to the vascular wallallowing the leukocytes to transiently stop. This activates the leukocytes by exposingthem to endothelial derived cytokines, resulting in up-regulation of the32-integrinsthat tighten adhesion and direct emigration (McEver, 1992). During inflammation,cell adhesion molecules are transiently expressed to recruit leukocytes. This isfollowed by the down-regulating of adhesive events, which limits and terminates the40recruitment of leukocytes. Firm adhesion and emigration of leukocytes throughactivated endothelium results in the shedding of L-selectin as a de-adhesion event(Kishimoto et aL, 1989; Porteu and Nathan, 1990; Spertim et aL, 1991b; Tedder,1991). One explanation proposed for this phenomenon is that chemotactic stimulationof leukocytes results in shedding of L-selectin from the cell surface (Julita et at,1989; Kishimoto et at, 1989; Griffin et al., 1990). Whether the process of rolling, thecontact of the PMN with chemotactic factors on the endothelium or the migrationprocess itself induce shedding is still unknown.Similar shedding of L-selectin occurs when lymphocytes cross high endothelialvenules (REV) in homing to peripheral lymphnodes (Kishimoto et at, 1990; Spertiniet at, 1991b). Notwithstanding the increase in information concerning the role of Lselectin in mediating leukocyte-endothelial cell interaction during physiologicalhoming of leukocytes to lymphnodes or recruitment to foci of inflammation, little isknown regarding adhesion events when leukocytes cross the endothelial barrier fromthe bone marrow to the circulating blood.In the bone marrow, L-selectin is expressed on nearly all postmitotic myeloid cells(Griffin et al., 1990), and a study by Lund-Johansen et al (Lund-Johansen andTerstappen, 1993) shows that L-selectin expression increases with granulocytematuration in the bone marrow with the highest expression on band cells andsegmented PMN in the bone marrow. However, this study did not directly comparethe expression of postmitotic bone marrow PMN with circulating PMN. The fact thatthe crossing of an endothelial barrier is associated with L-selectin shedding led us to41consider the possibility that a similar shedding of L-selectin occurs when PMN crossthe bone marrow-blood barrier.42WORKING HYPOnIHESISThe evidence that the migration of PMN through an endothelial barrier is associatedwith the shedding of L-selectin from their surface led to the working hypothesis thatL-selectin is shed from the surface of PMN as they cross from the bone marrowhematopoietic compartment into the circulating blood. Therefore, the studies in thefirst part of this thesis were designed to measure L-selectin expression on PMN in thehematopoietic and sinusoidal compartments of the bone marrow and in the peripheralblood during an episode of active bone marrow release.SPECIFIC AIMSThis working hypothesis was examined by pursuing the following specific aims:1) To determine the expression of L-selectin on mature PMN in the bone marrowhematopoietic tissue (hematopoietic compartment), the bone marrow venous sinusoids(sinusoidal compartment) and the circulating blood.2) To study the changes in L-selectin expression on PMN in these three compartmentsduring active bone marrow release of PMN and when bone marrow release issuppressed by hypothermia.3) To determine the effect of hypothermia on the shedding of L-selectin from PMN invitro.43Model of bone marrow release of PMNThe PMN released from the marginated pool come from organs such as the liver,spleen and lung with the lung being the major source of marginated PMN (Doerschukand English, 1991; Doerschuk and Allard, 1989; Muir et al., 1984). Previous studiesduring cardiopulmonary bypass (CPB) demonstrated an inithi fall in circulatingPMN that is attributed to the activation of the complement system (Chenoweth et aL,1981; Haslam et aL, 1980). This neutropenia is followed by a neutrophilia, whichmust be due to either a mobilization of marginated PMN from a pool other than thelung or to a release of new cells from the bone marrow. As the lung represents themajor pool of marginated PMN and as it is removed from the circulation during CPB,newly released PMN from the bone marrow form the largest component of theleukocytosis observed during CPB. An increase in fragments of C3 has been shownto occur minutes after commencing CPB (Chenoweth et al., 1981; Quiroga et al.,1985) and these fragments, such as C3e, have been associated with the bone marrowrelease of leukocytes (Alper et aL, 1972). Animal studies have shown that theaccumulation of PMN in the lungs of the animals after complement activation istwice as many as could be accounted for by the population of PMN in the circulation(Doerschuk and Allard, 1989). As the release of the PMN from the liver or spleen isunlikely in these circumstances (Doersehuk, and English, 1991; Rosolia et al., 1992),this suggests that PMN are released from the bone marrow following complementactivation. The leukocytosis associated with CPB-surgery has also been shown to betemperature dependent (Quiroga et al., 1985); that is the rise in circulating PMN canbe prevented by lowering body temperature to 27°C. This observation provides uswith an opportunity to study bone marrow release of PMN under conditions of active44release and when this release is suppressed by hypothermia. The fact that L-selectinis expressed on nearly all myeloid cells in the bone marrow (Griffm et aL, 1990) andthat removal of L-selectin from the cell surface is postulated to occur throughenzymatic cleavage by an unidentified cell associated protease that may betemperature sensitive (Spertini et al., 1991b; Tedder, 1991) led us to consider that thetemperature sensitive bone marrow release of PMN (Quiroga et al., 1985) is relatedto a reduction in cleavage of L-selectin from the PMN surface.45METHODSExperimental procedures:The goals outlined by the specific aims were pursued in studies on volunteers and onpatients who were undergoing normothermic and hypothermic CPB-surgery.In vitro studies:The effect of temperature on the shedding of L-selectin from the surface of PMN wasfirst determined in vitro using PMN isolated from the blood of 10 healthy infection-free laboratory workers. Zymosan activated plasma (ZAP) was used to activate PMNto simulate in vivo complement activation seen during CPB-surgery. ZAP wasprepared by incubating human plasma combined with zymosan A yeast (Z-4250,Sigma), 5mg/mi plasma. The mixture was incubated for 30 minutes at 37°C, spundown twice at 500 x g for 10 minutes, and used within 30 minutes. In 5 experiments,leukocyte rich plasma (LRP) was prepared from blood collected in acid citratedextrose (ACD) (1:5) as anti-coagulant and sedimented using high molecular weight(Mw 100,000-200,000: 4%) dextran, final dilution 1.9% dextran (Chemical DynamicsCoop., New Jersey) for 25 to 30 minutes. Leukocyte rich plasma samples wereincubated at 37, 27 or 4°C for 15 minutes, stimulated for 4 minutes with a 5%, 1 %,0.1 % or 0.01 % dilution of ZAP at the above mentioned temperatures and then thecells were fixed with 0.05 % glutaraldehyde prepared in 0.2M phosphate buffer pH 7.6.Kishimoto et al have shown that 4 mm of maximal stimulation of PMN causeshedding of >90 % of L-selectin from PMN and significantly upregulated CD 18(Kishimoto et al., 1989). Leukocyte rich plasma prepared from each specimen wasused to make cytospins by cytocentrifugation at 180 x g on a Cytospin 2 (Shandon46Lab Products, Cheshire, England) for 4 minutes onto 3-aminopropyl-tri-ethoxysilanecoated slides. Leukocyte rich plasma was diluted with PMN buffer 1:1 (138mM NaC1,27 mM KC1, 8. 1mM NaIl2P04.7H20, l.5nM KH2P04, 5mM glucose, pH 7.4) or furtherif necessary to obtain a single layer of cells on the slide. The leukocytes on the slideswere stained with the APAAP method (Cordell et al., 1984) using anti-Leu-8 (BectonDickinson, 1mm Systems, CA) as the primary antibody for the presence of L-selectinas described under immunocytochemical analysis. In 5 separate experiments,potassium ethylenediamine tetra-acetic acid (EDTA) blood (100d) was stimulatedwith the same concentrations of ZAP (5 %, 1 %, 0.1 % and 0.01 %) at the differenttemperatures (37, 27 and 4°C) for 4 mm. PMN L-selectin (using FITC conjugatedanti-Leu-8) and CD18 (using FITC anti-CD 18, Sigma Chem, St Louis, MO) expressionwas determined by flow cytometry as described under flow cytometric analysis. Thesestudies were done to determine whether the shedding of L-selectin under hypothermicconditions is PMN activation dependent.In vivo studies:Patient population:Ten patients (3 women and 7 men, mean age 63 years, range 43-76) were recruitedfrom patients admitted to St. Paul’s Hospital, Vancouver, B.C., for elective CPB foreither coronary arterial bypass and/or valvular replacement surgery. Five patientswere studied during hypothermic and 5 during normothermic CPB procedures.Approval from our institution’s (St. Paul’s Hospital and the University of BritishColumbia) human experimentation committee was obtained for the study andinformed consent was obtained from all patients prior to surgery.47Experimental protocolThe surgery was performed under general anaesthesia with endotracheal intubation.The anaesthetic drugs used were thiopental, diazepam, fentanyl, non-depolarizingmuscie-relaxants and oxygen-nitrous oxide enflurane or isoflurane mixture. Accessto the thoracic cavity was obtained via a median sternotomy and specimens of bonemarrow were obtained from the cut surface of the sternum using a small curette.Bovine heparin was given intravenously before the introduction of the venous andarterial cannulas using a dose that is required to lengthen the activated clotting timeto 450 seconds (Hemochron) and was repeated to keep the activated clotting time inthat range during the CPB procedure. No immune-modulating drugs such as steroidswere used before or during the surgery.The CPB apparatus consisted of a Sarns roller pump, a cardiotomy reservoir(Bentley® 220f) and a Cobe Optiflow II bubble oxygenator supplied with 98 % oxygenand 2 % carbon dioxide. The CPB circuit was primed with 5 % Ringers lactate exceptin one case where packed red cells mixed with a crystalloid solution were usedinstead. Pulmonary blood flow was reduced by diverting the venous return from theheart to the oxygenator pump in steps until complete CPB was achieved and restoredat the end of the procedure by increasing venous return to the right heart stepwise.Systemic hypothermia was used in 5 patients with a perfusate temperature of 26-28°C. Patients were cooled until bladder temperature (core temperature) was 27°Cand rewarmed to 37°C at the end of the CPB procedure. In 5 patients, normothermiawas maintained by keeping the perfusate at 37°C.48Blood and bone marrow samplesPeripheral blood samples were obtained from an arterial line and kept at thetemperature at which they were collected. In preliminary experiments, samples weretaken across the CPB apparatus at different time points during bypass, to determinethe effect of PMN circulating through the bypass apparatus on PMN L-selectinexpression. No notable effect of the apparatus on PMN L-selectin expression werefound. These samples were collected: before the start of anaesthesia and surgery(baseline), after the sternum was split (BM1); before restoring pulmonary blood flowin the normothermic group and just before rewarming in the hypothermic group(BM2); and at the end of the surgical procedure just before the sternum was closed(BM3).Bone marrow samples were obtained from the sternum at time points BM 1, BM2 andBM3. Smears were made immediately and allowed to air dry. Bone marrow sampleswere processed for histology by embedding in glycolmethacrylate (GMA) using amodification of methods described for processing bone marrow tissue in plastic forimmunohistochemical analysis (Burgio et al., 1990; Murray and Evens, 1991). Plasticembedding of tissue is frequently use as an alternative to wax embedding for lightmicroscopical examination and GMA is the plastic of choice. Embedding the tissuein plastic had a variety of advantages in our study; thinner sections can be cut withimproved resolution of cellular detail, tissue shrinkage is less and tissue can beembedded without decalcification (in preliminary experiments we demonstrated thatdecalcification of bone marrow by either hydrochloric acid or EDTA rendered thetissue useless for detecting our antigen of interest).49As indicated above, bone marrow specimens were collected from the cut surface of thesternum with a small curette and immediately fixed in 0.025 % glutaraldehydeprepared with 0.2M phosphate buffer pH 7.6 for two hours at the temperature atwhich the tissue was collected. Fixation with this very low concentration ofglutaraldehyde has been shown in preliminary experiments to preserve morphologyand to have the least influence on the detection of our antigen of interest. Afterfixation, specimens were transferred to acetone at 4°C for 12 hours. The specimenswere then transferred to GMA resin monomer (JB4®; Polyscience, Ltd.) containing0.9% benzoyl peroxide-solution A and infiltrated under vacuum (-75mmHg) using aFreeze dryer Model FDU 101, Blazer Union, for 2 hours at 4°C. The infiltrated tissuewas placed in embedding molds and GMA chemically polymerized by a mixture of200L of JB4® solution B to 5m1 of solution A (JB4®; Polyscience, Ltd.) for 12 hoursat 4°C. Polymerized resin blocks were brought to room temperature and 2m sectionscut with a Sorval® JB4 microtome fitted with a glass knife (made with a LKB Knifemaker Type 7801B, Stockholm, Sweden). Sections were floated on a room temperaturewater bath, transfered to glass slides and air-dried overnight at room temperature toinsure adhesion of sections to the glass slides during the staining procedure.Hematological analysis:Peripheral blood samples were collected in standard tubes (Vacutainer, BectonDickinson, NJ) containing potassium EDTA. Blood cell counts and differential whitecell counts were performed with a Sysmex Model E4000 (Toa, Japan). Band cellswere identified (College of American Pathologists, 1991) and counted on Wrightsstained blood smears from all samples. The blood smears were evaluated in a blinded50fashion by evaluating 100 PMN in randomly selected fields. All leukocyte countswere corrected for hemodilution by means of the hematocrit;Corrected cell count = Observed cell count boseline hematocritobserved hematocritL-selectin determination:a) Evaluation of L-selectin expression on PMN in the circulating blondAll blood specimens were kept in an incubator and processed at the temperature atwhich they specimen was collected. Leukocyte rich plasma (LRP) was prepared fromblood collected in acid citrate dextrose as an anti-coagulant and cytospun on coatedslides as has been previously described. Slides were air-dried and stained within 7days. All cytospins prepared from the peripheral blood were fixed in acetone for 10minutes prior to immunocytochemical staining.1) Immunocytochemical staining for L-selectin:The alkaline phosphatase anti-alkaline phosphatase (APAAP) technique was used todetect the presence of L-selectin on cells (Cordell et al., 1984). Specimens wereincubated with 5 % rabbit serum for 15 mm to block non-specific binding of thesecondary linking antibody. This incubation was followed by the application of theprimary antibody anti-Leu-8 (5g/ml) at room temperature in a humidity chamber for60 minutes. All antibodies were prepared in Tris Buffered Saline (TBS) pH 7.6 for 60minutes and ±301il of the antibody solution was used per slide. Antibody dilutionswere determined empirically by preliminary titration experiments and found to be51saturation at 5gIm1. Non-immune mouse IgG (5 Jhg/ml) and the omitting of theprimary antibody were used as negative controls. As a linking antibody a 1/20dilution of rabbit anti-mouse IgG (Z259 DAKO, Denemark) was applied for 45minutes; and finally, the anti-mouse conjugated with alkaline phosphatase complexes(D651 DAKO, Denemark) in a 1/50 dilution was applied for 45 minutes. Slides werewashed in TBS for 5 to 10 minutes following each antibody application. The alkalinephosphatase was developed by a final incubation of 20 minutes in a substratecontaining 50mg naphthol-AS-BI-phosphate, in 0.6ml N-N-dimethylformamide addedto 0.5m1 of 4% sodium nitrite solution with 0.2m1 fuschin (Merck 4040: 5g in lOOmi2M HCL) and lOOmi of TBS pH 8.7. Endogenous alkaline phosphatase was blockedby adding 17.5mg levamisole to the solution. The slides were counterstained withfiltered Harris-hematoxylin for 60 seconds. All slides were mounted in an aqueousmounting medium and evaluated on a Zeiss Universal Research light microscope(Model 1W, West Germany) at 400X magnification.2) Immunofluorescent flow cytometryImmunofluorescent staining ofcirculating PMN was done to determine the expressionof L-selectin with FITC-conjugated anti-Leu-8 (Becton Dickinson, CA) using flowcytometry. A whole blood method for preparing specimens was used. The cells wereprepared from EDTA blood using a commercially available kit (Coulter Clone®,Coulter Electronics, Florida). Briefly, 100d of EDTA blood was incubated with 2001.dPBS buffer and 0.24g FITC-conjugated anti-Leu-8 for 10 mm at room temperaturein the dark. For each blood sample, a negative control was done using non-immunemouse FITC-conjugated IgG2a (Becton Dickinson, CA). After washing the cells twice,the red blood cells were lysed (Immuno-lyse, Coulter Clone®). The remaining52leukocytes were fixed with 1 % paraformaldehyde and stored at 4°C. Flow cytometrywas performed on the specimens within 24 hours (Model; Profile EPIC II, CoulterElectronics, Florida). Analysis gates for the PMN were established using thedistinctive forward and side scatter profiles. A total of 3000 gated cells wereevaluated per specimen and are presented as either percent positive cells, meanfluorescence intensity (log), or histograms.b) Evaluating L-selectin expression on bone marrow specimens1) Bone marrow smears were made when bone marrow samples were collected, air-dried and stored at room temperature, and stained within 7 days using the methoddescribed for cytospin specimens.2) Immunohistochemical staining of plastic embedded sections of bone marrow for thepresence of L-selectin was performed by a three layered biotin-avidin alkalinephosphatase technique. Sections were allowed to hydrate for 15 minutes in 0.5M Trisbuffered saline, pH 7.6 (Sigma Chemical Co, St.Louis, MO), trypsinized with asolution of 0.05% trypsin (Sigma Chemical Co, St Louis, Mo) prepared in TBS at 37°Cfor 5 minutes. Endogenous avidin binding activity was blocked by successive 20minute incubations of sections with avidin 0.1 % and 0.01 % biotin (Dako, Denemark)(Woods, and Wanrke, 1981). The monoclonal antibody anti-Leu-8 (Becton Dickinson,CA) was used to label L-selectin. Each section was incubated for an hour at roomtemperature in a humidity chamber with lOOd of a 0.5g/ml solution of anti-Leu-8prepared with 1 % BSA TBS. Non-immune mouse IgG (Sigma Chemical Co, St.Louis,MO) was used in a similar dilution as the primary antibody as negative control.Omitting the primary antibody was done as an additional control. Slides were53incubated for 30 minutes in biotin conjugated goat anti-mouse IgG (Fc receptorspecific) 15g/ml. A goat serum blocking step was omitted as goat sera do not reactwith Fe receptors of human leukocytes (Alexander and Sanders, 1977). Slides wereincubated for 30 minutes with avidin (4gIm1) conjugated with alkaline phosphatase(DAKO, Denemark). All slides were washed twice in TBS for 15 mm between eachantibody application. The alkaline phosphatase was developed for 15 minutes in thedark with a commercially available kit (Histomark® Red, Kirkegaard and Perry Lab,Gaithersburg, MD). This is a new Fuchsin-naphthol AS-BI phosphate systemproducing a red-scarlet reaction product. The sections were washed with TBS,counterstained with Mayer’s-hematoxylin for 60 seconds, air-dried and mounted withCoverbond®.Immunocyto- and histochemical grading systemsOn the cytospin and bone marrow smears, PMN and band cells were evaluatedaccording to the intensity of staining of the cell surface using an arbitrarily designedgrading system, grading PMN from negative (GO) to highly positive (deep red stainingof more than 75 % of the PMN surface area, G4) (figure 4a-f). The slides were codedand examined without knowledge of their origin from either the normothermic orhypothermic CPB group. Fields were selected in a systematic randomized fashionand 100 cells were evaluated per specimen. All cells of interest in a selected fieldwere evaluated except if the cell was broken or overlapping with other cells, asoverlapping cells tend to trap stain between them. The inter-observer variability wasdetermined by having two observers grade 10 randomly selected slides, and the intraobserver variability by having one observer repeating the measurements 3 weeks54later without knowledge of the origin of the slides or the initial results.The intensity of staining of PMN on the histological specimens of bone marrow wasless than on the cytological specimens (cytospins and BM-smears); therefore, PMNwere graded from negative or background (GO) to highly positive (deep red stainingof more than 50 % of the PMN surface, G3). Inter-observer variability of the gradingsystem was tested by having 3 observers evaluate 100 cells from a total of 15photomicrographs randomly selected from 10 patients and the intra-observervariability by one observer evaluate the same cells 3 weeks apart without knowledgeof the initial results. The reproducibility of the visual grading system was furthertested against a computerized high resolution true colour (1024 x1024, 24bit) imageanalysis system (BioviewsInfraScan®) used in our laboratory that determines theintensity, saturation and hue of a colour in an area that is digitized. One hunderdrandomly selected PMN were evaluated for the fraction of surface area that stainedintense red and were compared to the visual grading system in a blinded fashion.Quantitative histologyThe L-selectin expression on PMN in sinusoids and in the hematopoietic tissue of thebone marrow were quantified using standard morphometric techniques (Cruz-Oriveand Weibel, 1981; Gundersen, 1977; Steno, 1984). Toluidine Blue 0 (TBO) was usedto delineate the vascular structures in the bone marrow. Immunocytochemicalstaining of bone marrow endothelial cells (factor 8 and CD3 1) was used to confirm thepresence of vessels in the bone marrow identified by TBO. However, both of thesevessel markers resulted in background staining of PMN that made doubleimmunocytochemical labelling of sections difficult to interpret. Therefore, L-selectin55Figure 4: Photomicrographs (A) to CE) represent the range of staining of PMN in thecirculation. PMN were graded according to the intensity of staining from: (A) GO(negative): (B) Gi (positive andJor <25% deep red stain of the cytoplasm): (C) G2(between >25 and <50% ofdeep red stain of the cytoplasm): (D) G3 (between >50 and<75% of deep red stain of the cytoplasm): to CE) G4 (highly positive, more than > 75%deep red stain of the cytoplasm). Cells excluded were broken cells, overlapping cellsand mononuclear cells. Both circulating PMN prepared from leukocyte rich plasmaand bone marrow smears were stained by the APAAP method (see text).Photomicrograph (F) demonstrates the range of staining of PMN typically seen in asingle field (from highly stained cells to negative cells). The bars represent a lengthof 5gm.56on PMN and the vasculature of the bone marrow were stained on two adjacent serialsections. The number of PMN per unit volume of bone marrow was estimated usingthe disector method (Steno, 1984). Five 2m thick serial sections of bone marrowwere cut and labelled. Sections two (reference section) and four (lookup section) (4gmapart) were stained for the presence of L-selectin, and sections number one and fivewith TBO to delineate the vasculature. Section three was stained with hematoxylinand eosin and used to identified cells and structures between slides two and fourwhich were used as the disectors (figure 5). In five randomly selected fields bonemarrow sinusoids were identify in section one. The same sinusoidal vessel wasidentified in sections 2, 4 and 5 and all areas of interest photographed with a ZeissUniversal research lightmicroscope (Model 1W, West Germany) on 35mm Kodak®Gold (ASA 100) film and printed as 100 x 150mm photographs with a finalmagnification of photographs of 3,630X. Vessels were delineated on photographs ofsections 2 and 4 (figure 6b) using photographs of sections 1 and 5 (figure 6a). Volumefractions of hematopoietic tissue, vessels, fat spaces and bone trabeculae wereestimated by using standard point counting techniques with quadrilateral unbiasedcounting chambers placed over corresponding areas on photographs of section 2 and4 using landmarks such as bone spicules and megakaryocytes (Cruz-Orive, andWeibel, 1981; Gundersen, 1977). All PMN as well as band cells within the chamberin the reference photograph (section 2) that did not appear in the lookup photograph(section 4) were marked. Mature PMN were identified by their unique nuclear shape,dense chromatin with no nucleoli and granular cytoplasm if visible. All marked PMNwere graded according to their intensity of staining as mentioned previously. Section1, 3 and 5 were used to verify the nature of any doubtful cells. Photographs wereSectionSerial sections for Disector5.71 TBO2 L-selectin < Reference34Spare (H&E)L-selectin Look-up5 TBOFigure 5: The disector method to quantify the number of PMN in each bone marrowcompartment. Five 2m thick serial sections were cut and numbered 1-5. Sections 1and 5 were stained with Toluidine Blue 0 (TBO) to delineate the vasculature andsection 2 (reference section) and 4 (lookup section) for the presence of L-selectih.Section number 3 was stained with hematoxylin and eosin and used to identify cellsand structures between slides 2 and 4.Figure 6: Photomicrograph (a) shows a bone marrow sinusoid demonstrated with aToluidine Blue 0 stain of a 2m thick section of bone marrow embedded inglycolmethacrylate and photomicrograph (b) the corresponding serial section stainedfor the presence of L-selectin using the avidin-biotin alkaline phosphatase technique(see text) on PMN (arrow). The broken line represents the vascular margin of thesinusoid in the reference section. These PMN were graded from negative (GO) tohighly stained (G3) when >50% of the PMN surface area stained deep red. The barsrepresent a length of 10im.5S’59evaluated in a single blinded fashion without knowledge of the patient from whichthey came. A calculated total of 0.03 135mm of bone marrow tissue was examinedfrom each specimen. This was calculated:VBM = Lx Wx Dwhere VEM represents the total volume of bone marrow examined for each specimen,L the length, W the width and D the depth of the disector. The length and widthmeasured on the microphotograph were converted to real length by dividing by thefinal magnification of the tissue on the microphotographs. The number of PMN perunit volume (lxmm3) of bone marrow sinusoids and hematopoietic tissue wascalculated. For example the number of PMN in the sinusoids was calculated:N=Vv x Vsinusoid BMwhere N,vSflUSQd is the number of PMN in the sinusoids of a particular specimen perone cubic millimetre of bone marrow, the number of PMN counted in thespecimen, the volume fraction of sinusoids in the specimen and VBM thevolume of bone marrow examined in each specimen. Similar calculations were donefor the hematopoietic tissue and the number of cells in each grading category. All theresults were expressed as the number of PMN in each grade per cubic micrometer ofbone marrow compartment.60Statistical analysis:Statistical analysis was performed using SYSTAT® Version 5.1 software (Systat, Inc.,Evanston, IL) (Wilkinson, 1990).The corrected white blood cell, PMN and band cell counts at each time point werecompared between groups and over time using a generalized linear model forrepeated measurements. Differences in L-selectin expression on PMN after ZAPstimulation were evaluated using a two-way analysis of variance for eachtemperature. Differences between PMN L-selectin expression on circulating versusbone marrow cells and differences between time points within groups and betweengroups were analyzed using a two-way analysis of variance with Bonferronicorrections and testing for multiple comparisons. With data presented as apercentage of baseline (MFI), differences between experimental time points weretested for statistical significance by comparing 95 % confidence intervals for differenttimes. Individual time points were considered statistically different from baseline ifa 95 % confidence interval for that time point did not include 100%. Students t-testwas used to compare demographic variables between groups. A probability of lessthan 0.05 was considered to be significant. The data are expressed asmean± standard error except when otherwise mentioned.Immunocyto- and histochemical scoring systemsInter- and intra-observer variations in the immunocytochemical grading system wereevaluated by calculating the scoring differences noted by two observers or by oneobserver grading slides 3 weeks apart without knowledge of the identity of slides.Differences within the range of the 95% confidence intervals (mean±2xSE) were61considered to represent acceptable reproducibility. The inter- and intra-observervariation of the immunohistochemical scoring system were evaluated by calculatingthe Pearson coefficient of mean-square contingency(R 2) for each grade and expressingR2 as a fraction of the maximum possible value, R2 (Sachs, 1982). The Pearsoncoefficient of mean-square contingency is an extension of the Pearson chi-square test(the null hypothesis being that the rows and columns of a matrix are independent).In the immunohistochemical grading score, R2max would represent the value of thePearson chi-square coefficient if there is 100% agreement in the grading scorebetween and within observers: thus R= ±0.82. For analysis of both the inter- andthe intra-observer variation, an R2IR ratio >0.75 was considered to representacceptable reproducibility. A Spearman rank correlation was used to compare thevisual grading system with the computerized image analyzer.62RESULTS:Surgical procedure:Table II shows data concerning the patient population studied and variables in thesurgical procedure. Although the total duration of the surgical procedure was longer(p<O.Ol) in the normothermic group, the time between the collection of BM1 andBM2 and the pump times were not significantly different between the two groups.The longer surgical time was because more of the patients in the normothermic grouprequired combined procedures (coronary bypass and valve replacements).Hematological results:The total white blood cell count, PMN cell count and band cell count rose fromthe beginning (BMI) to the end (BM3) of CPB surgery in both groups (Table Ill).These data also show that the increase in white cell, PMN and the peripheral bloodband cell counts observed between BM1 and BM2 in the normothermic group(p <0.01) was prevented by hypothermia. Rewarming the patient after the completionof surgery was associated with a rise in both the percentage of penpheral blood PMNand band forms to levels that is comparable to the length of the surgery. The absolutenumber of these cells were lower than in the normothermic group, which wasconsistent with the total duration of the procedure.Immunocytochemical grading system:Examples of the staining achieved for L-selectin in cytological specimens usingimmunocytochemistry are shown in figure 4a-f. The inter- and intra-observerdifferences were less than 5 % for grading both negative (GO) and highly positive PMN‘3TABLE IICharacteristics of Cardiopulmonary Bypass PatientsNormothermic Hypothermicn=5 n=5Age (years) 67.2±8.3 57.4±10.8Sex (MJF) 4/1 1/4Surgery:Coronary Bypass 3 2Valve Replacement 0 3Both 2 0Pump time (mm) 120±27 68±26Time cooled at 27°C 0 21±13.5Surgery time (mm) 160±36 93±55*with paired T-test64Table ifiPeripheral leukocyte counts during Cardiopulmonary Bypass surgeryNormothermic Hypothermicn=5 n=5Corrected White Cell Count:Baseline 6.9±0.48 4.8±0.51BM1 6.4±0.59 4.7±0.42BM2 16.7±3 3.1±0.81BM3 21.8±3.25’ 11.8±4Corrected PMN Cell Count:Baseline 4.9±1.52 2.9±0.33BM1 4.6±0.47* 2.8±0.29BM2 13.1±2.12 1.9±0.65BM3 18.7±2.2’ 9.1±3.15Corrected Band Cell Count:Baseline 0.25±0.07 0.26±0.09BM1 0.18±0.03* 0.15±0.04BM2 2.98±0.56 0.59±0.28BM3 6.24±1.06’ 2.67±0.83All leukocyte counts were corrected for hemodilution using the hematocrit andexpressed as number x109/L. Baseline, BM1, BM2 and BM3 are time points whenblood was collected (see text).Note that hypothermia prevented the rise in the white cell, PMN and band cellcounts that occurred in the normothermic patients between BM1 and BM2. Thesecounts was partially restored when the patients were rewarmed to normothermia atthe end of surgery (BM3).Difference between BM1 and BM2 in the normothermic group, <0.05.Difference between baseline and BM3 in the normothermic group (‘p <0.01) andhypothermic group ($p <0.05).65(G4). Because intra-observer variation in grading the intensity of staining of PMNwas the smallest for the negative (3 ± 2.1, mean ± 2xSE) and highly positive (grade 4+)cells (4± 2.9), these observation grades were used by preference when comparingdifferences between time points and groups (figure 7). Similar results were obtainedin histological sections with all the R2/R values >0.75 with the highest valuebeing in the GO grade (0.91). There was a good correlation between the visual gradingsystem and the computerized image analyzer (Infra-Scan) grading PMN (R=O.9)(figure 8).ImmunocytochemistryTable IV shows that the segmented PMN and band cells in the bone marrow stainedstrongly for L-selectin with the majority of these cells faffing in the G4 category.Under baseline conditions the peripheral blood PMN stained less intensely in bothgroups with more GO and Gi cells and less G4 cells (p< 0.01) (figure 9). Undernormothermic conditions when there was bone marrow release, this differencebetween blood and bone marrow become smaller (see table IV). However, underhypothermic conditions, where bone marrow release was slowed, the difference in GOand G4 cells between blood and bone marrow persisted (p<O.Ol) (table IV). Figure 10demonstrates the changes in PMN L-selectin during cardiopulmonary bypass (BM1to BM2). Over this time period (BM1 to BM2), the number of highly positive PMN(G4)increased significantly in the circulation of the normothermic group (9±3.3 to36 ± 6.6 %, p <0.03) but not in the hypothermic group(10 .7±4.7 to 18 ±5.7%, p =NS)(table IV). The expression of L-selectin on band cells was marginally lower than onsegmented PMN in the bone marrow smears (figure 1 la) but significantly higher40-1. 1I I I— I I0 GO Gi G2 G3 G4Figure 7: Variability in grading the expression of L-selectin on PMN in cytologicalspecimens. Slides were coded and examined without knowledge of their origin and inrandomized fields on each slide a 100 segmented and non-segmented PMN weregraded (see grading system figure 4). Grades are expressed as a fraction of the 100cells counted. The graph demonstrates the inter-observer variability when oneobserver graded 10 slides 3 weeks apart without knowledge of the origin of the slidesor the initial results. The y-axis represents the mean difference between the twoobservations and the error bars are two standard errors. The variability in gradingnegative (GO) and highly positive (G4) cells was small (<5%).()ci)ci)‘4-‘4-U27141-12-25IGrade‘7100-R=’0.9180C). 60-Cct1.... 0C)40- HDCl)0- 20-‘4-C0 I I I IGO Gi G2 G3 G4Visual grading systemFigure 8: The correlation between the visual and Infrascan® grading systems. A 100PMN on five randomly selected slides. Cellsd were stained (red) for the presence ofL-selectjn with the APAAP technique. PMN were graded from negative (GO) to highlystained (G4) visually (see grading system, figure 4) and compared to thecomputerized high resolution true color image analysis system determining the areaof the cell surface that stained intense red (y-axis is area of intense red). There wasa good correlation (Spearman rank correlation, R=O.91) between the two gradingsystems.Go-.•)aSFigure 9: The difference between L-selectin expression on PMN in the circulatingblood (panel a) and the bone marrow (bone marrow smears, panel b). In thecirculating blood a wide range of staining intensity (GO to G4) was seen in contrastto the bone marrow where the majority of PMN were highly stained (G4) (see tableIV). Bar is 1OtmTable IVChanges in PMN L-selectin expression during cardiopulmonary bypassGrades GO Gi G2 G3 G4Normothermic:BM1: Blood* 26.2±6.5 37.4±6.4 17.8±2.5 9.4±2.4 9±3.3tBone marrow 0.4±0.2 1.8±0.9 14.6±2.6 35±3.4 47±4.3BM2: Blood 8.2±5.2 12.8±2.68 16±3.1 27.4±3.6 36±6Bone marrow 0.7±0.4 5.1±1.6 18.5±1.5 31±4.2 45±2.7Blood 4.4±2.4 12.2±3.6 18.8±4 31.8±3 32.6±6.4Bone marrow 1.7±1.4 4.5±1.5 18.3±5.1 26.5±3.6 49±8.3Ilypothermic:BM1: Blood* 28±10.1 27.2±3.7 17.7±3.1 15.7±4 10.7±4.7Bone marrow 0.2±0.1 4.2±1.1 15.8±4.3 22.6±3 57.4±6.3Bloods 17.8±11 16.8±4.2 18.6±3.6 29.4±8 18±5.7Bone marrow 0.4±0.4 9.6±1.7 21.2±2 26.4±5.9 42.2±6BM3: Blood 8.2±4.8 18.6±6.4 16±1.9 24.2±4.2 32.6±8.6Bone marrow 1.4±1.4 7.2±3.3 15.4±2.1 28.6±3 47.4±4*p <0.01, $p <0.05; Difference between L-selectin expression on bone marrow and circulating PMN.tp < 0.03; An significant increase in highly stained PMN with bone marrow release in thenormothermic group (BM1 to BM2) which was suppressed in the hypothermic group.Grades; On bone marrow smears and PMN in circulating blood, a 100 PMN were graded fromnegative (GO) to highly positive (G4), see text for grading system (figure 4).BM1,BM2 and BM3 referred to time points blood and bone marrow specimens were collected, see text.‘ci70z0CC.)BM1 BM2Figure 10: Immunocytochemical determined L-selectin expression on circulating PMNat the start (BM1 before bypass, solid bars) and the end (BM2 end of bypass, hatchedbars) of normothermic cardiopulmonary bypass surgery (a). With bone marrow releaseof PMN in the circulation the number of strongly positive (G4) cells increased andnegative (GO) cells decreased (*p <0.01). However, when active bone marrow releasewas suppressed by hypothermia (b), L-selectin expression on circulating PMNremaines the same. One hundred PMN were counted for each specimen and the y-axisrepresents the fraction of those PMN. Values are the mean±SE of 5 patients.a*LJbGO Gi 02 G3 04Grades71than segmented PMN in the circulation (figure 1 lb).Flow cytometry:Figure 12 is an example of the changes in L-selectin expression on circulatingPMN as measured by flow cytometry during normothermic (figure 12a-c) andhypothermic (figure 12d-f) cardiopulmonary bypass. Figure 13 shows the dataobtained by flow cytometry where the expression of L-selectin (Mean FluorescenceIntensity, MFI) on circulating PMN in the normothermic group increased in astepwise fashion from 10.8±4.4 at baseline to 12.2±4.9. This represents a 16±9.8%increase in MFI(log) of circulating PMN L-selectin expression at the end of the bypassprocedure (p <0.05). In the hypothermic group, on the other hand, the MFI changedlittle over the same period and tended to fail at the end of the surgical procedure.Morphometric studies:The staining intensity of PMN in the histological specimens of bone marrow was lessthan that achieved on bone marrow smears due to the light glutaraldehyde fixationstep required to preserve morphology and the processing of the tissue intoglycolmethacrylate. In each group one patient was excluded from the morphometricanalysis because of inadequate (hypoplastic) bone marrow tissue. At baseline in bothgroups (figure 14), a significant difference in the expression of L-selectin on PMN wasseen between the sinusoids and hematopoietic compartments with the fraction ofnegative PMN the highest in the sinusoids (24.7±3.5 versus 10.3 ±2.5 %, p <0.004)and L-selectin positive PMN (G1-G3) higher in the hematopoietic compartment(88 ± 3.1 versus 69.5 ±4.8%, p <0.005). The expression of L-selectin on PMN in the72.‘-4CCUF70 -a605040CC: 30U2O10-0——GO G4bTGi G2 G3GradesI_WA0GOSegm PMNG2 G3 G4GradesGiBandsFigure 11: The expression of L-selectin on segmented PMN (solid bars) and band cells(hatched bars) in the bone marrow (a) and the circulating blood (b). Note the smalldifference in L-selectin expression between segmented PMN and band cells in thebone marrow but a significant difference between these cells in the circulation duringbone marrow release of PMN. Slides were randomly selected from time points (BM2and BM3) with peripheral band cell counts >10% of total PMN counts, allowing 100band cells to be evaluated per slide. Values are the mean±SE of 13 time points.73Mean Fluorescence IntensityFigure 12: Expression of L-selectin on PMN during bone marrow release of PMN ina normothermic (panels a to c) and hypothermic (panels d to f) patient as measuredby flow cytometry. Note the increase in L-selectin expression in the normothermicpatient during bone marrow release of PMN (BM1 to BM3). The two populations ofcells seen in panel (b) may represent a population of PMN that have shed their Lselectin with intravascular cell activation (left) and a population of PMN releasedfrom the marrow with high levels of L-selectin (right). No evidence of an increase inL-selectin during bypass was seen in the hypothermic patient (BM1 to BM2). In eachspecimen 3000 PMN were evaluated and expressed as mean fluorescence intensityon a Log scale (x-axis).BM1 BM2a2i.._ :d..BM3D0C.)G)C)Cb2A e—a2f.: 1—74N=5Figure 13: Expression of L-selectin on circulating PMN as measured by flowcytometry at the different time intervals in 5 normothermic and 5 hypothermic CPBprocedures. In each specimen 3000 PMN were evaluated and expressed as meanfluorescence intensity (MFI) on a Log scale. Values in this graph are expressed asa percentage of baseline MFI values (obtained before anaesthesia and surgery start).All values are the mean±SE of 5 specimens. In the normothermic group thepercentage changes in the MFI over time increased significantly (*p <0.05) whereashypothermia was associated with either no change or a slight decrease in MFI at theend of surgery.Norm=normothermic groupHypo =hypothermic group140124108CtJ 9276*- NormHypo60Baseline BM1 BM2 BM3Time Intervals75hematopoietic compartment did not change over the study period (Table V). Figure15 shows that with bone marrow release of PMN in the normothermic group, the Lselectin negative PMN (GO) increased from 5.75±1.1 x 10 at baseline (BM 1) to14.6±2.8 x iO PMN/mm3 at the end of bypass (BM2), P <0.03, whereas in thehypothermic group it stayed the same (6.2 ± 1.2 to 6.3 ±0.82 x iO, p =NS). Thisincrease in the GO PMN in the sinusoids in the normothermic group represents anincrease from 24 to 46 % of all the PMN in the sinusoids (Table V). These datademonstrate a gradient of PMN L-selectin from the hematopoietic tissue to thesinusoids at baseline that increased during active bone marrow release of PMN andremained the same when marrow release was prevented by hypothermia.In vitro studies of L-selectin:Figure 16 shows the flow cytometry data concerning the expression of L-selectin andCD18 on PMN incubated at 37, 27 and 4°C stimulated with increasing concentrationsof ZAP ranging from 0 to 5%. At 37°C, PMN shed L-selectin and upregulated CD18in a dose dependent manner.Incubation of the cells at 27°C (figure 1 6b) suppressedthe effect of ZAP stimulation except at the highest dose and incubation at 4°C (figure16c) eliminated the effect of ZAP even at the highest dose. PMN incubated at 27°Cand stimulated with low concentrations of ZAP (0.01 and 0.1 %) were activated asreflected by the increased CD18 expression. However, little changes in PMN Lselectin expression was seen at these low concentrations of ZAP stimulation. Thisdissociation of CD18 upregulation and L-selectin shedding was not apparent withPMN incubated at 37°C. These findings were supported by immunocytochemicalstaining and grading of PMN on cytospins of peripheral blood, which showed no76za00C,I.IL0Hematopoietic SinusoidsFigure 14: Combined baseline data for both normothermic and hypothermic groups.The fraction of cells negative for L-selectin (GO) was greater in the bone marrowsinusoids than in the bone marrow hematopoietic tissue in both groups of patients(*p <0.004), and the fraction of positive PMN (sum of Gi, G2 and G3) was greater inthe hematopoietic compartment (‘p < 0.005). Values are the mean±SE with n=8.100-80-60-40-20-*#GOG1G2G3TABLEVL-selectinexpressiononPMNinthebonemarrowVenoussinusoidsHematopoietictissueGradeG0G,G2G3G0S,G2G3NormoBM15750109922927350047633873339926832115329592164197418291480125514365BM214623*131411597241571042254688509504289617399328773247259158885640BM362381542957334366534284571074616329342172757334365258309175036939Hypo BM162171429436714748522935404102691314612324081153220871293490913233501BM26330120235159635249612779911471232078282049201421085855294116712286BM36761149557636660273033463558801784881516534175391731561108220848362ThevaluesrepresentsthenumberofPMN/mm3ineachbonemarrowcompartment.Thevaluesinitalicsarethestandarderrorofthemeanoffourpatients.*RepresentsasignificantincreaseofGOgradedPMNfromBM1toBM2(p<O.03).7g$.—00zwzFigure 15: Changes in the number of negative graded PMN (GO) in the marrowsinusoids at the start (BM1) and at the end (BM2) of bypass. With bone marrowrelease of PMN in the normothermic group (hatched bars), the number of GO PMNincreased significantly (p < 0.03). However, when bone marrow release was preventedby hypothermia, the number of GO cells in the sinusoids remained the same. The yaxis represents the number of PMN per cubic millimetre of bone marrow. Values arethe mean±SE of four patients in each group.*p<003 *2010-0BM1HypothermicB2____NormothermicFigure 16: Flow cytometry data concerning the expression of L-selectin and CD18 onPMN incubated at 37, 27 and 4°C in vitro and stimulated with increasingconcentrations of zymosan activated plasma (ZAP) ranging from 0 to 5%. The solidbars represent L-selectin and the hatched bars CD18 expression. Note that at 37°C,PMN shed L-selectin and upregulate CD18 in a dose-dependent manner. However,these changes are suppressed at 27°C and abolished at 4°C. At 27°C a dissociationbetween L-selectin shedding and CD18 upregulation was seen with stimulation ofPMN with low concentrations (0.01 and 0.1%) of ZAP. Values of mean fluorescenceintensity are the mean±SE of 5 experiments with all values expressed as a fractionof baseline.*p<O.05$p <0.027’?37°C1% 5%27°C *T0%300-0.01% 0.1%4-I0C)4-IC)0C)C)0C)I0LIciC)*240-180-120-80-00.01% 0.1% 1% 5%4°C0% 0.O% 0.1% 1%Zymosan Activated Plasma80change in the number of negative PMN when stimulated with low concentrations ofZAP at 27°C in contrast to a significant increase at 37°C (figure 17).Figure 17: Changes in the negative graded PMN (GO) incubated at 37, 27 and 4°Cstimulated with no(control or 0%), 0.1% and 5% zymosan activated plasma. PMN oncytospin specimens were stained for L-selectin by the APAAP method in 5experiments. Note the significant increase of L-selectin negative (GO) cells with amild stimulus (0.1% ZAP) in the 37°C incubated cells that was not seen in the 27°Cincubated cells.*p<o.05#p<O.05*z0ci)>C)C4-’C)a)C)08I4°C27°C— 37°CP010IT0% 0.1% 5%Zymosan Activated Plasma82DISCUSSIONThis study confirms several previous reports showing that there is systemicleukocytosis during CPB-surgery (Chenoweth et al., 1981; Quiroga et aL, 1985) andthat hypothermia prevents this leukocytosis by suppressing the bone marrow releaseof PMN. The quantitative histology of the bone marrow and the in vitro studies linka temperature sensitive L-selectin loss from the surface of the PMN to a temperaturesensitive release of these cells from the bone marrow hematopoietic tissue into themarrow sinusiods. They further demonstrate a PMN L-selectin gradient between thebone marrow and the circulating blood that decreases with active bone marrowrelease of PMN.Evidence for bone marrow release of PMN:The PMN storage pool in the bone marrow is estimated to be 5.6 x iO cells/kg with± 40 % of these cells being mature segmented neutrophils and ± 60 % being band formsand metamyelocytes (Boggs, 1967). Under stable conditions, the PMN released frombone marrow are mainly mature segmented PMN and these cells form the bulk of theso-called marginated pool of cells. The marginated cells can be rapidly mobilized intothe circulation with moderate stress such as exercise (Muir et al., 1984; Foster et al.,1986) whereas more severe stress associated with trauma, infection or complementactivation is required to release band cells and rarely metamyelocytes into thecirculation. Although the ratio of segmented to band cells released from the bonemarrow is not known and probably varies with the type and the intensity of thestimulus, an increase in the circulating band cell count provides definitive evidencethat PMN are being released from the bone marrow. Our study shows an increase83in circulating PMN during CPB-surgery and the source for this increase is fromeither the intravascular marginated or the bone marrow pools. As bypass of thepulmonary circulation removes a major source of marginated cells (Peters et aL, 1985)the release of cells from the bone marrow is the most likely source of the increasednumber of circulating leukocytes in these studies. Furthermore, release from themarginated pool is characterized by an increase in circulating PMN without aconcomitant increase in the number of circulating band cells (Boggs, 1967;Hetherington and Quie, 1985). Release from the bone marrow, on the other hand, ischaracterized by an increase in the number of circulating band cells in the peripheralblood (Boggs, 1967; Hetherington and Quie, 1985; Marsh et aL, 1967; Jagels andHugli, 1992). In this study, the band cells increased from 5 % of the total PMN countat baseline to more than 25 % of the circulating pool at the end of surgery in bothgroups (see table Ill). The fact that hypothermia suppressed the total number of bandcells in the circulation and rewarming restored band cell counts to normothermiclevels confirms the previous report from our laboratory that the bone marrow releaseof PMN is a temperature dependent phenomenon.Evidence for L-selectin loss when PMN cross the bone marrow-blood barrierThe release of bone marrow stores of PMN appear to largely accounts for theperipheral neutrophilia induced by many factors known to promote leukocytosis(Athens et al., 1961; Kajita and Hugh, 1990; Jagels and Hugli, 1992) includingintravascular complement activation. Chemotactic factors that induce neutrophiliacause morphologic and adherent properties of the stimulated cells that are short-lived. It seems unlikely that PMN recruitment from the bone marrow represents a84chemotactic response per Se, because a gradient would exist for only a brief period ifat all. However, stimulation of bone marrow cells by chemotactic factors may induceboth morphologic and cytoskeletal changes within the PMN, as well as modificationsand reorganization of cell surface molecules involved in adherence between PMN andeither endothelium (Griffin et al., 1990; Von Andrian et al., 1991) or stromalelements of the bone marrow.L-selectin is expressed by nearly all circulating PMN more or less continuouslythroughout myeloid differentiation in the bone marrow (Griffin et al., 1990).Lewinsohn et al (1987) demonstrated that in mice mature bone marrow PMNexpressed higher levels of L-selectin (MEL-l4) than peripheral circulating PMN, butwe know of no previous quantitative study comparing the expression of L-selectin onPMN in the bone marrow compartments of humans. Our data show that theexpression of L-selectin was highest on PMN in the hematopoietic compartment(figure 14) in the bone marrow and that this compartment did not change to asignificant extent over the study period (see Table V). PMN in the bone marrowsinusoids expressed less L-selectin than those in the hematopoietic compartment withmore L-selectin negative and less L-selectin positive PMN. The baseline differencein L-selectin expression between the PMN in the sinusoids and those in thehematopoietic compartment (figure 14) suggests that L-selectin is shed when crossingthe bone marrow-blood endothelial barrier or gradually lost as PMN are releasedfrom sinusoids into the circulation. Our observation that there is a difference in PMNL-selectin expression across the bone marrow- blood barrier at baseline could haveseveral explanations. The immature myeloid cells in the bone marrow make close85contact with the processes of adventitial reticular cells and may be anchored to thesecells or the matrix through pro-adhesive lectin-like adhesion molecules. Gradual lossof these molecules during maturation under stable conditions might permitmovement towards the sinus walls and eventual release into the circulation.Alternatively, the process of crossing the endothelial barrier may result in a loss ofL-selectin from the PMN surface similar to that which occurs with recruitment ofPMN into an area of inflammation. A third possibility is that the mature PMN andband forms that have entered the marrow microvasculature are anchored to themarrow endothelium by L-selectin with subsequent cleavage resulting in a rapidentry into the circulation with an overshoot leukocytosis such as that seen incardiopulmonary bypass surgery.L-selectin is expressed on granulocytes and is known to shed from the surface of PMNduring cell activation (Smith et al., 1991; Tedder, 1991). Although the mechanismand regulation of shedding is unknown, enzymatic cleavage of the surface receptorby a membrane bound protease with chymotrypsin activity is the most likelypossibility (Spertini et aL, 1991b; Tedder, 1991). Cell activation is thought to inducechanges in the conformation of the L-selectin protein and expose nascent sites thatare susceptible to enzymatic cleavage. The fact that the release of PMN from thebone marrow during rewarlning after CPB occurs over a narrow temperature rangefrom 35-37°C (Quiroga et al., 1985) led us to consider the possible role of atemperature sensitive enzymatic cleavage of L-selectin in the release of PMN fromthe bone marrow. Our data demonstrate an increase in the number and fraction ofL-selectin negative PMN in the bone marrow venous sinusoids with active bone86marrow release of PMN (figure 15 and Table V) which was also accompanied by anincrease in the total number of PMN in the sinusoids (Table V). This suggests thatthe leukocytosis induced by complement activation not only released PMN thesinusoids but also involves egress of PMN from the hematopoietic compartment to thevenous sinusoids.This increase was not seen under conditions where the bone marrow release of PMNwas suppressed. The in vitro studies suggest that the activity of the enzymeresponsible for L-selectin cleavage from the activated PMN surface was suppressedby reducing the temperature to levels that prevent marrow release. This reductionin L-selectin shedding was independent of PMN activation by ZAP stimulus becauseCD18 was upregulated when L-selectin shedding was suppressed at 27°C. Thisfunctional dissociation between PMN activation and L-selectin shedding suggests thatthe enzyme responsible for L-selectin cleavage is functionally disabled at 27°C andtotally inactive at 4°C. It also suggests that the enzyme responsible for the cleavageof L-selectin during marrow release may not be one of the granular proteases releasedfrom activated cells.The concept that bone marrow release is related to the loss of L-selectin is supportedby the previously reported observation that myeloid cells with low expression of Lselectin tend to be released from the bone marrow into the circulation (Griffin et al.,1990). It is also of interest that patients receiving GM-CSF develop marrowhyperplasia with an increased number of circulating progenitor cells (Platzer, 1989)and that GM-CSF induces shedding of L-selectin from all myeloid cells including the87progenitor cells. These observations are consistent with the hypothesis that cleavageof L-selectin from the cell surface contributes to the relocation of progenitor cells fromthe marrow into the blood (Griffin et al., 1990). Furthermore, GM-CSF has beenshown to induce complete shedding of L-selectin from circulating PMN within onehour of administration, with low L-selectin expression on the PMN persisting for upto 6 days (Demetri and Antman, 1992). This implies that the resulting leukocytosisfollowing GM-CSF administration may be related to cleavage of L-selectin frommyeloid cells in the bone marrow. Spertini et al (1991b) have demonstrated that acutemyelomonocytic leukemia and chronic myeloid leukemia cells rarely express Lselectin, in contrast to their normal counterparts. The lack of L-selectin expressionon these cells could explain their relocation from the bone marrow into thebloodstream.The present studies show that shedding of L-selectin from the surface of PMN can beinhibited by hypothermia, both in vivo and in vitro. They also show that underbaseline conditions, the expression of L-selectin is greatest on the PMN in thehematopoietic compartment of the bone marrow and that this expression is decreasedin the bone marrow sinusoids. Although several possible mechanisms could explainthis result, the data clearly show that L-selectin expression is reduced as the PMNleave the marrow and that this change is prevented when marrow release issuppressed by hypothermia.Regulation of cell-matrix and cell-cell interactions via the expression of adhesionpromoting receptors on the surface of leukocytes has been proposed as a possible88mechanism for controffing the egress of PMN from bone marrow (Campbell, andWicha, 1988; Hynes, 1992; Miyake et a!., 1990a; Simmons et aL, 1992 Springer,1990b; Williams et al., 1991). An example of this mechanism is that the loss ofreceptors for the universal attachment peptide, arginine-glycine-aspartic acid (RDG),on erythroid progenitors is believed to initiate the release of reticulocytes from themarrow (Tsai et aL, 1987). Another adhesion protein in the bone marrowextracellular matrix, hemonectin, binds more specifically to myeloid cells and mayanchor them in the marrow until full maturation (Campbell et al., 1987a). However,the nature of this protein, the receptor on the myeloid cells for hemonectin and thefactors that control this adhesion event are not known. In this study we have linkedthe release of PMN from the bone marrow to the shedding of L-selectin from theirsurface. This adds to the body of evidence that the adhesion promoting molecules playa critical role in the egress of leukocytes from the bone marrow. We speculate thatshedding of L-selectin from the surface of PMN may be an important step in theirrelease from the bone marrow.Evidence of increase in expression of L-selectin on circulating PMN during active bonemarrow releaseL-selectin is expressed by nearly all circulating PMN and post-mitotic myeloid cellsin the bone marrow (Collins et al., 1991; Griffin et al., 1990; Spertim et al., 199 ib).Lewinsohn et a! (1987) demonstrated in mice that mature bone marrow PMNexpressed higher levels of L-selectin (MEL- 14) than peripheral circulating PMN, witha reduced expression of L-selectin (IVIEL-14) on more immature myeloid cells(Lewinsohn et al., 1987). Lund-Johansen et al (1993) have also recently demonstrated89in that humans L-selectin increases on bone marrow PMN with cell maturation. Weknow of no other study directly comparing the expression of L-selectin on PMN in theperipheral blood and bone marrow compartments of humans during active bonemarrow release of PMN.Evaluating bone marrow smears showed at baseline the expression of L-selectin washigher on bone marrow PMN than on cirrculating PMN in both the normothermicand hypothermic groups and that it did not change to a significant extent over thestudy period (Table IV). There was less L-selectin on circulating PMN (more negativeor Gi cells and fewer highly positive or G4 cells) than on the bone marrow PMN inboth groups of patients under baseline conditions (Table IV). Furthermore, as thePMN on bone marrow smears are diluted by blood from the bone marrow venoussinusoids, our estimate of the L-selectin expression on PMN in the hematopoieticcompartments is probably low, and the PMN L-selectin gradient between thehematopoietic compartment and the circulating blood is probably even larger thanobserved. The difference in L-selectin expression between bone marrow PMN andPMN in the circulation under baseline conditions suggests that L-selectin may be lostas PMN cross the bone marrow-blood barrier or that circulating PMN progressivelylose L-selectin during their intravascular life. As bone marrow smears contain amixture of cells from the hematopoietic tissue and bone marrow venous sinusoids, thereduction in PMN L-selectin crossing the bone marrow-blood barrier is unlikely to bethe sole reason for the difference in L-selectin expression we observed between bonemarrow smears and cytospins. Loss of L-selectin from the PMN during their lifespanin the general circulation is a likely explanation and this hypothesis will be explored90in the second part of this thesis.L-selectin expression on circulating PMN increased and approached the level ofexpression on bone marrow cells during normothermic CPB surgery, but it alwaysremained lower than on bone marrow PMN (Table IV). As this increase occurredwhen the PMN were being released from the bone marrow, it is consistent with newcells containing more L-selectin entering the circulation. The increase in peripheralblood PMN expressing the highest levels of L-selectin (G4) during bone marrowrelease in the normothermic group supports this concept. The increase in PMN Lselectin expression in the circulation during bone marrow release of PMN suggeststhat notwithstanding the lost of L-selectin in the bone marrow during PMN release,the released PMN expression of L-selectin is still higher than PMN in the circulationunder baseline conditions. This may imply that the bone marrow-blood barrierbecame more permeable during active release with less shedding of L-selectincrossing the barrier. The bone marrow-blood barrier has been described as becomingmore permeable during active bone marrow release of PMN (Weiss, 1970, Tavasolli.,1977). The alternate possibility that PMN upregulate their L-selectin expressionwhile they are in the circulation is much less likely because there are no reports ofeither in vitro or in vivo upregulation of L-selectin in mature PMN during cellactivation. Furthermore, the low levels of L-selectin mRNA in circulating cellssuggest either message instability or low levels of translation and processing (Tedderet al., 1989) and imply that L-selectin is unlikely to regenerate on circulating PMN.When the bone marrow release of PMN was suppressed by hypothermia, the L91selectin expression on circulating PMN remained close to baseline levels (figure lOb).However, restoration of bone marrow release by rewarming the patient resulted inan increase in the number of peripheral blood PMN expressing high levels of Lselectin. These observations establish that the level of L-selectin expression on thecirculating PMN is highly dependent on their release from the bone marrow.Assuming that the majority of band cells in the circulation have been recentlyreleased from the bone marrow, our finding that band cells in the circulation expresshigher levels of L-selectin than segmented PMN (figure 1 ib), supports the hypothesisthat peripheral blood PMN expressing high levels of L-selectin have been recentlyreleased from the bone marrow.Interestingly, the expression of L-selectin on band cells in the bone marrow was lowerthan on segmented PMN, which suggests that the mature segmented PMN in thebone marrow contain the most L-selectin. This observation supports the findings ofLund-Johansen et al (1993) that L-selectin expression increases with PMN maturationin the bone marrow. The segmented mature PMN in the bone marrow have beenshown to be more deformable and reactive to chemotactic stimuli than immaturemyeloid cells (Lichtman, 1970). These are the cells released from the bone marrowunder baseline conditions (Rosolia et al., 1992; Ulich et al., 1989). With a stimulusfor bone marrow release of PMN, the mature PMN are the first to enter thecirculation. With a more intense or prolonged stimulus, band cells and rarelymetamyelocytes appear in the circulation (Deinard and Page, 1974; Ghebrehiwet andMuller-Eberhard, 1979; Kampschmith, 1984; Platzer, 1989). As such a stimulus alsoactivates intravascular PMN to shed L-selectin, our finding of an increase in L92selectin on circulating PMN shows that the cumulative effect of complementactivation during CPB surgery favoured bone marrow release.The significantly lower L-selectin expression on circulating PMN than bone marrowPMN can be partly explained by the loss of PMN L-selectin when crossing the bonemarrow-blood barrier. Notwithstanding this loss, L-selectin on circulating PMNincreased with bone marrow release of cells. Therefore, the wide distribution of Lselectin expression on circulating PMN can be explained as a mixture of cells withhigh L-selectin expression that have recently been released from the bone marrowand a population of older PMN that have lost some or all of their L-selectin whilethey are in the circulation. Cardiopulmonary bypass results in bone marrow releaseof PMN expressing high levels of L-selectin, which increases the population ofcirculating PMN expressing high levels of this cell adhesion molecule.Summary1) We have shown that under baseline conditions the expression of L-selectin isgreatest on bone marrow PMN and that there is a stepwise decrease in expressionfrom the hematopoietic compartment in the bone marrow to the bone marrowsinusoids, and a further decrease to the circulation.2) The shedding of L-selectin from the surface of PMN can be inhibited byhypothermia, both in vivo and in vitro.3) Our data further support the concept that L-selectin is partly shed from the PMNas they cross from the hematopoietic compartment into the bone marrow venoussinusoids with a stimulus for bone marrow release such as complement activation.934) With bone marrow release of leukocytes, L-selectin expression on circulating PMNincreased, supporting the concept that circulating PMN expressing high levels of Lselectin are cells recently released from the bone marrow.945) CHANGES IN L-SELECTIN ON CIRCULATING PMNIntroductionThe selectin family of adhesion-promoting molecules, including L-, E- and P-selectin,are found on endothelial cells (E- and P-selectin), platelets (P-selectin) and leukocytes(L-selectin). They mediate leukocyte rolling and margination of PMN in post capillaryor collecting venules (Ley et al., 1993; Ley et aL, 1991b; Von Andnan et aL, 1992;Von Andrian et aL, 1991), the earliest manifestation of leukocyte recruitment. Lselectin has been shown to be essential in the leukocyte-endothelial interactioncascade of events in vitro and in vivo that result in PMN recruitment to foci ofinflammation (Butcher, 1991; Ley et aL, 1991b; Mulligan et aL, 1994; Von Andrianet al., 1991; Springer, 1 990b). Recruitment of PMN to the inflamed peritoneal cavityof mice can be blocked by blocking antibodies to L-selectin or L-selectin-Ig chimera(Jutila et aL, 1989a; Jutila et al., 1989b; Watson et al., 1991). In two models of PMNdependent acute lung injury in rats (intrapulmonary deposition of Ig-G complexes andintravascular complement activation), the requirement of L-selectin has beendocumented (Mulligan et aL, 1994). These data suggested that L-selectin plays animportant role in the recruitment of PMN during the acute inflammatory reactionin the systemic and pulmonary vessels.The selectin-dependent initial adhesive interaction between the leukocytes andendothelium is followed by the second phase of leukocyte recruitment, which appearsto depend on the engagement of the f32-integrins (CD 11/CD 18) on the leukocytes aswell as ICAM-1 and ICAM-2 on the endothelium for the firm adhesive interactionthat accompanies eventual transmigration into the extravascular space (Butcher,951991; Larson and Springer, 1990; Zimmerman et al., 1992). This transmigration eventhas been shown to result in the loss of L-selectin from the surface of the PMN(Kishimoto et al., 1989; Julita et al., 1989). However, if the conditions for firmadhesion and emigration are inadequate, PMN may return to the circulation.Whether this latter event results in a loss of PMN L-selectin is unknown.This hypothesis is attractive considering:1) Shedding of L-selectin could provide a rapid means for the regulation of leukocyteadhesion to the endothelium with the subsequent release of PMN back into thecirculation. Receptor-ligand binding would initiate receptor shedding and serve as ameans of receptor modulation. This concept is attractive considering the fact that justa small fraction of PMN delivered to a focus of inflammation eventually emigrate(Coxson et al., 1990; Doerschuk et al., 1994).2) The interaction of leukocytes with “normal” unactivated endothelium may alsoresult in L-selectin shedding. Using intravital microscopy, leukocytes tumble and“roll” on unactivated endotheium, which becomes more pronounced when theendothelium are activated (Lawrence, and Springer, 1991; Smith et aL, 1991). In thepulmonary capillary bed, leukocytes make intimate contact with the endothelium andhave to deform to negotiate their passage through the lung. Both of these events mayresult in PMN L-selectin being lost. The demonstration of high levels of free Lslectin in plasma under normal conditions suggests that leukocytes continuouslyshed L-selectin in the circulation (Schleiffenbaum et al., 1992).3) The variable expression of L-selectin on PMN in the circulation with populationsof cells expressing high, intermediate and low levels of L-selectin contrasts with thePMN released in the circulation from the bone marrow which expresses high levels96of Lse1ectin (as dicussed in previous chapter). This suggests that L-selectin is lostfrom the surface of PMN during their lifespan in the circulation.The second part of this thesis explores the fate of L-selectin on PMN during theirlifespan in the circulation.97WORKING HYPOTHESISOur working hypothesis is that:1) Circulating PMN shed L-selectin from their surface during their normalintravascular life resulting in “older” PMN expressing lower levels of L-selectin.2) Low L-selectin expression on “older” PMN is a signal for removal of these cellsfrom the intravascular pool.As L-selectin is an activation sensitive cell surface receptor, this hypothesis had tobe tested using PMN labelled and transfused to recipients with the least possiblemanipulation.SPECIFIC AIMSThis hypothesis was examined by pursuing the following specific aims:1) To develop a method of labelling PMN in vivo that decreases or eliminates theneed for in vitro purification and subsequent labelling of PMN, all steps that maypotentially activate PMN. To accomplish this goal an animal model was developedin which PMN were labelled in vivo with the thymidine analogue 5’bromo-2’-deoxyuridine (BrdU).2) To establish the best method for transferring labelled PMN to recipients in ouranimal model and to determine the clearance of these labelled PMN from thecirculation as well as their functional capabilities.N3) To transfuse labelled PMN to recipient animals and to determine the expressionof L-selectin on these labelled PMN over time in the circulation.4) To determine whether the lack of L-selectin expression on circulating PMN servesas a signal for the removal of these cells from the intravascular pool.98Model of in vivo PMN labelling:The following specific aims will be addressed in this section:1) To develop a method of labelling PMN in vivo that decreases or eliminates theneed for in vitro purification and subsequent labelling of PMN, all steps that maypotentially activate PMN. To accomplish this goal an animal model was developedin which PMN were labelled in vivo with 5’bromo-2’-deoxyuridine (BrdU).2) In this model, to establish the best method for transferring labelled PMN torecipients and to determine the clearance of these labelled PMN from the circulationas well as their functional capabilities.Previous studies from this (Doerschuk et al., 1987a; Muir et al., 1984) and severalother laboratories (Segal et al., 1976) have reported on the in vivo behavior of PMNlabelled with isotopes such as chromium (51Cr),indium (“1n), ordiisopropylfluorophosphate (DP32P). These isotope studies require the PMN to beisolated from whole blood, labelled with an isotope in vitro and transfered back intorecipients. These procedures make the experimental protocol both lengthy andexpensive (Haslett et al., 1985; McAfee et a!., 1984) with subsequent decontaminationand disposal of radioactive waste. There is also the possibility that the purificationand labelling procedures will activate the PMN and cause them to behave abnormally(Haslett et al., 1985). The classical method for detecting cells engaged in DNAsynthesis in vivo is by their uptake of the thymidine analogues ([3H1-thymidine, [3H1-TdR), identified using autoradiography (Denekamp and Kaliman, 1973). Theincorporation of[3H]-TdR allows in vivo labelling of cells but provides few if any99advantages over the in vitro labelling with gamma emitting isotopes (Denekamp andKallman, 1973; Hamilton and Dobbin, 1983). However, the thymidine analogue,BrdU, which was labelled with the radioisotope 82bromine, in early studies (Hakala,1958; Eidnoff et aL, 1959), has now been modified for immunocytochemical detectionusing a monoclonal antibody with high specificity against this halogenated analogue(De Fazio et al., 1987b; De Fazio et a!., 1988; Dolbeare et aL, 1985; Gratzner, 1982).The immunocytochemical visualization of BrdU appears to be a powerful alternativeto[3H]-thymidine autoradiography and this technique has been used increasingly tomonitor cell proliferation and migration during normal development (Dombrowicz etal., 1988; Plickert et al., 1988), DNA replication (Gratzner, 1982), particularly instudies involving growth kinetics in solid tumors (Denekamp and Kallman, 1973;Goodson et al., 1991; Soriano and Del Rio, 1991), and cytofluorometric analysis of thecell cycle (De Fazio et al., 1987b; De Fazio et al., 1988; Dolbeare et al., 1983). Theimmunocytochemical staining is performed within hours compared withautoradiography which take days and often weeks. The normal PMN turnover isbetween 100 and 200 billion cells a day (Dancey et al., 1976; Walker and Willemze,1980) and labelling these rapidly dividing cells in the bone marrow with BrdU mayprove to be ideal.By using BrdU as a non-isotopic method to label PMN in vivo, the behavior of thesecells can then be studied by transfusing whole blood to serum compatible recipients.This method may eliminates the need for in vitro purification and subsequentlabelling of PMN, all steps that may potentially activate PMN. It will further allowsus to study the behavior of an activation sensitive surface molecule such as L-selectin100over time in the circulation. Furthermore, it may also reduces the cost of disposalof isotopes and isotope contaminated specimens and animals.Described here is a method for labeffing and detecting the proliferating pool of PMNin the bone marrow by administering BrdU to donor animals. The behavior of theselabelled cells was then studied in serum- compatible recipients after the cells hadbeen transferred in either whole blood, leukocyte rich plasma, or purified PMN.101METHODSAnimals:Twenty female New Zealand white rabbits were used in this study. Three (4 ±0.3 kg, mean ± SD) were used as donors and 17 (2.6 ± 0.2 kg) as recipients. Thestudy was approved by the Animal Experimentation Committee of the University ofBritish Columbia.Labelling DNA of rabbit leukocytes with BrdU:Donor rabbits were given BrdU (Sigma Chemical Co., St. Louis, MO) at a dose of 25mg/kg daily for 7 days. The BrdU was infused slowly through the marginal ear veinat a concentration of 5 mg/mi in normal saline over a period of 15 minutes. Bloodobtained daily from the central ear artery of these BrdU-treated rabbits was analyzedas follows: lml was collected in standard Vacutainer tubes containing potassiumethylene diamine tetra-acetic acid (Becton Dickinson, Rutherford, NJ) for blood cellcounts determined on a model SS8O Coulter Counter (Coulter Electronics, Hialeah,FA) and for differential white cell counts on Wright’s stained blood smears; and 2m1was collected in acid-citrate-dextrose (ACD) for the preparation of leukocyte richplasma (LRP) as previously described. The resulting LRP was cytospun at 180 x gfor 4 minutes to obtain a monolayer of cells on pre-coated slides (Fisher Scientific Co.,Pittsburgh, PA), air dried, and stained to determine the number of BrdU labelledPMN in each specimen. After labelling efficiency of PMN was evaluated, theserabbits were used as a source of labelled PMN in all subsequent experiments.102PMN purification:PMN were purified from 25 ml of donor blood as previously described from ourlaboratory (Doerschuk et aL, 1987a; Doerschuk and English, 1991). Briefly, LRPobtained from 25 ml of donor blood was centrifuged and suspend in 1 ml PMN bufferand hypotonic lysis of the residual red blood cells in the LRP was achieved bydilution in 11 ml sterile water. After 18 seconds, 1 lml 2X PBS (27 mM Na2HPO,132mM KH2PO4, and 2.74 M NaC1) and 10 ml PMN buffer were added. PMN wereseparated from the mononuclear cells by centrifugation in Histopaque (SigmaChemical Co.) with a density of 1.077 g/ml at 150 x g for 13 minutes. The isolatedPMN were 95 % to 98 % pure with a viability of 97% as assessed by trypan blueexclusion.Transfer of BrdU labelled leukocytes to recipient rabbits:BrdU labelled leukocytes were transferred from donor to recipient rabbits as either25 ml whole blood, LRP prepared from 25 ml blood, or PMN purified from 25 mldonor blood. Cells were transfused into the marginal ear vein of serum-compatiblerecipient over a 5 mm period. The total number of leukocytes, PMN, BrdU labelledPMN, and the amount of BrdU labelled DNA in the 3 preparations was determinedfrom fractions of each donor sample. Eight recipients received whole blood, 3 receivedLRP, and 6 recieved purified PMN. The behavior of BrdU labelled PMN in thecirculating blood was assessed from blood samples obtained from the central earartery of all recipients before the BrdU labelled PMN were infused (baseline) at 5, 30mm, and at 1, 3, 6, and 24 hours. The blood samples were used to determine whiteblood cell and differential cell counts, the percentage of BrdU labelled PMN in103cytospun preparations of LRP, and the fraction of BrdU labelled DNA.Assessing the number of BrdU labelled PMN in the circulation:The percentage of BrdU labelled PMN in the peripheral blood of recipients wasdetermined by counting a minimum of 200 PMN on each cytospin slide on randomlyselected fields. If less than 5 % of PMN were labelled, 500 PMN were counted, if lessthan 1 % of PMN were labelled a 1000 PMN were counted. The number of BrdUlabelled PMN in the circulation of each recipient was expressed as a fraction of thetotal number of labelled PMN originally infused and was corrected for the calculatedblood volume (American Physiology Society, 1965) of the recipient in the followingmanner:BrdU BV x %PMNrB.JFraction PMN. =_______________________Cirt BrdUedwhere Fraction PMNU represents the number of BrdU labelled PMN in thecirculation as a fraction of the total number of BrdU labelled PMN infused, PMNthe calculated number of PMN (1 x 106) in the circulation (total white cell count timesthe fraction of leukocytes that are PMN) times the calculated blood volume(BV),%PMN the fraction of BrdU labelled PMN in a cytospin of peripheral blood inthe recipient, and PMN the number (1 x 106) of BrdU labelled PMN infused(PMN count/mi x ml of fluid infused x %BrdU labelled PMN). In three rabbits whoreceived whole blood more frequent blood samples were taken: at baseline, 2.5, 5, 7.5,10, 15, 20, 30, and 45 minutes and 1, 2, 3, 4, 5 and 6 hours after infusion. Twenty-104four hours after infusion of the labelled cells, recipients were anaesthetized withintramuscular injections of 100 to 150 mg/kg of ketamine hydrochloride (Ayerst,Philadelphia, PA) and 5 to 8 mg/kg acepromazine maleate (Ayerst) and sacrificed ina supine position by an injection of 2 to 3 mI/kg of saturated KC1 into the aortic root.Specimens of lung, liver, spleen, bone marrow, and gut were removed for theimmunohistochemical detection of BrdU labelled cells and for the analysis of BrdUlabelled DNA.Tmmunocyto- and histochemical detection of BrdU labelled cells:The lung, spleen, liver, and gut specimens were fixed in 10 % buffered formalin for2 to 4 hours while the bone marrow specimens was fixed with B5 fixative. Randomlyselected blocks from the fixed tissues were embedded in paraffin. Three m tissuesections placed on slides coated with 3-aminopropyl-triethoxysilane (Sigma ChemicalCo.) were baked for 16 hours at 37°C. The paraffin was removed in two 10 minutewashes of xylene. For sections of bone marrow the second xylene wash contained 4 %iodine in order to remove mercury remaining from the B5 fixative. Sections wererehydrated in graded ethanol from 100% to 70 %, rinsed twice with distilled water,and digested at 37°C for 10 minutes in a 0.4% pepsin (Sigma) solution acidified to pH2.5. All cytospun preparations of LRP were fixed in methanol for 10 minutes. DNAin both the tissue sections and cytospun specimens was denatured in 2 N HC1 at 37°Cfor 1 hour. This was followed by neutralization in three washes of 0.1 M boratebuffer, pH 8.5, each for 10 minutes. The APAAP technique (Cordell et al., 1984) wasused to detect BrdU labelled DNA in cells. Briefly, specimens were incubatedconsecutively in 5 % rabbit serum for 15 minutes, then in 0.5 g/ml mouse anti-BrdU105antibody (Boehringer-Mannheim, Mannheim, Germany) prepared with 1 % BSA in 50mM Tris Cl, 150 mM NaC1, pH 7.6 (TBS), at room temperature in a humidifiedchamber for 1 hour. Non-immune mouse IgG at 0.5g/ml was used as a negativecontrol in addition to a negative control where the primary antibody was omitted.Incubation in a 1/20 dilution of rabbit anti-mouse IgG (DAKO Laboratories,Copenhagen, Denmark) for 45 minutes was followed by 45 minutes in a 1/50 dilutionof a mouse monoclonal alkaline phosphatase anti-alkaline phosphatase complex(DAKO Laboratories). Slides were washed in 0.1 % Tween 20 in TBS for 10 minutesfollowing each antibody application. The alkaline phosphatase was developed for 20minutes in 100 ml TBS at pH 8.7, after the addition of a mixture of 0.5 ml of 4%sodium nitrite, 0.2 ml of 5 % fuschin (Merck, Rahway, NJ) in 2M HC1 and 50 mgnaphthol-AS-BI-phosphate (Sigma Chemical Co.) dissolved in 0.6 mlN,N-dimethylformamide. Endogenous alkaline phosphatase was blocked by theaddition of 17.5 mg levamisole (Sigma Chemical Co.) to the colour reaction. Thepreparations were counterstained with Mayer’s-hematoxylin for 60 seconds, mountedin an aqueous medium, and analyzed on a Zeiss Model flR Universal Research lightmicroscope (Oberkochen, Germany).DNA Analysis:Samples of individual organs were frozen in liquid nitrogen and stored at -70°C.Nuclei were isolated from whole blood, LRP, and PMN according to the method ofBuffone and Darlington (Buffone and Darlington, 1985) and DNA was isolated fromthese nuclei as well as from the lung, liver, spleen, skin, bone marrow, and ileumfollowing procedures described previously (Strauss, 1990). Ten g of DNA isolated106from blood, LRP, or PMN and 20g of DNA isolated from the above mentioned organswere electrophoresed through 0.7% agarose gels. The DNA was transferred from thegels to Hybond-N filters (Amersham, Arlington Heights, IL) according tomanufcturer’s instructions. After cross-linking the DNA to the ifiters withultraviolet light at 254 nm for 4 minutes, BrdU labelled DNA was detected usingmouse anti-BrdU antibodies following the APAAP method as described above. Thesefilters were washed for 5 minutes in 10 mM Tris Cl, 1 mM EDTA, pH 8.0,photographed, and analyzed on a Ultroscan XL densitometer using the Gelscan XLsoftware (Phamacia, Uppsala, Sweden).Morphometry:A stereologic analysis to determine the distribution of BrdU positive PMN (PMNB1J)in the lung, spleen, liver, gut and bone marrow tissue was performed on paraffinembedded tissue sections that had been immunologically stained for the presence ofBrdU. A point counting technique (Cruz-Orive and Weibel, 1981) was applied usinga Nikon microscope with a camera-lucida attachment, which allowed random fieldsof view generated by a computer program to be analyzed at 400X magnification bymeans of a point counting grid of 400 points that was superimposed onto themicroscope image. A total of 20 fields were evaluated per organ. The volume fraction(V) of BrdU labelled PMN was calculated from the equation:V = E poinLs )flPMN&dUV&W) E on the gridThen, the total number of PMNBrdU in each organ was calculated from the equation:107organ volume xNumber of PMN&dU = (PMN140 m3where PMNBrdU represents the number of BrdU labelled PMN and 140 m3 representsthe assumed volume of a PMN (Doerschuk et aL, 1993). The calculated total numberof pjBrdU in each organ was expressed as a percentage of the total number ofPMNU infused into each recipient rabbit.Activation parameters on BrdU labelled PMN:a) Expression of L-selectin and CD18 on BrdU labelled PMNImmunofluorescence staining of labelled and control PMN was done to determine theexpression of L-selectin (Monoclonal antibody DREG 200, kind donation of Dr. E.C.Butcher) and CD18 (Monoclonal antibody 60.3, kind donation of Dr. J.M. Harlan)using flow cytometry. Changes in the expression of these molecules on PMN aresensitive parameters of cell activation (Kishimoto et al., 1989; Julita et al., 1991;Julita et aL, 1989). A whole blood method for preparing specimens was used. Thecells were prepared from EDTA blood for analysis using a commercially available kit(Coulter Clone®, Coulter Electronics, Florida). Briefly, 100ILl of EDTA blood wasincubated with 2001.d PBS buffer and 0.4g MoAb DREG 200 or 60.3 for 10 mm atroom temperature in the dark. For each blood sample, a negative control was doneusing non-immune mouse IgG (Sigma Chem Co, St Louis, MO). After washing thecells with PBS buffer, FITC goat anti-mouse 0.24g was added to each specimen andincubated for 10 mm. After washing the cells twice, the red blood cells were lysed108(Immuno-lyse, Coulter Clone®). The remaining leukocytes were fixed with 1 %paraformaldehyde and stored at 4°C. Flow cytometry was performed on thespecimens within 24 hours (Model; Profile EPIC II, Coulter Electronics, Florida).Analysis gates for the PMN were established using the distinctive forward and sidescatter profiles. A total of 3000 gated cells were evaluated per specimen and arepresented as either percent positive cells or mean fluorescence intensity (log).b) Superoxide production of BrdU labelled PMN:The possibility that PMN are primed by the labelling procedure was evaluated bydetermining PMN superoxide anion production with N-formyl-L-methionyl-L-leucyl-Lphenylalanine (FMLP) and phorbol 12-myristate-13-acetate (PMA) (Sigma, St Louis,Mo) stimulation on purified labelled and control PMN, using the superoxidedismutase (SOD) inhibitable reduction of ferricytochrome C as described by Markertet al (1984). We preincubated 2x106/ml PMN for 5 mm at 37°C with and without23g/ml SOD. To initiate the reaction we added ferricytochrome C (80MM), and theappropriate stimulant, either PMA (2nM) or FMLP (10M). All reagents were finalconcentrations in a final volume of 750l and were incubated at 37°C for 30 mm. Thereaction was ended by adding 400jl isotonic Ranks buffered saline (HESS), pH 7.4,to the mixture and centrifuging it for 3 mm at 2000 x g. We measured the increasein absorbance at 550mm in a Perkin Elmer Lamda2 UV/VIS Spectrometer andconverted the results to nanomoles of reduced cytochrome C by using the extinctioncoefficient 18.5 mM’ .cm1 Values of superoxide produce by a final PMN concentration1 x 106 cells over 30 mm were determined by calculating the difference betweensamples with and without SOD. Each determination was done in duplicate and109expressed as nanomoles 02/3OminIlxlO 6 PMN.c) Migration of BrdU labelled cells into inflammatory fociA focal pneumonia was produced as has been previously described (Doerschuk et aL,1990) to test the ability of labelled PMN to migrate into a focus of infection. Briefly,5 x iO S. pneumoniae organisms were dissolved in 0.5 ml sterile saline containing5 % colloidal carbon. This mixture was instilled into the left lower lobe of twoanaesthetized rabbits through a paediatric feeding tube inserted between the trachealrings and positioned under fluoroscopy. An equal volume of sterile saline with 5 %colloidal carbon was instilled into the right lower lobe to serve as a control. Two 1cm x 1 cm foam sponges, one soaked by immersion in the solution of S. pneumoniaeand the other in the control solution, were then inserted subcutaneously on oppositesides of the linea alba. The animals were maintained anaesthetized in the proneposition for 3 hours by the administration of additional ketamine HC1 as required.At this point, 25 ml whole blood in ACD from a BrdU labelled donor was transfusedintravenously as above and the rabbits were sacrificed 1 hour later. Specimens ofpneumonic and control lung areas marked by the colloidal carbon and of skinsurrounding the sponges were excised for immunohistochemical detection of BrdUlabelled cells and for analysis of BrdU labelled DNA.STATISTICAL METHODSAll values are expressed as mean± standard error when except otherwise mentioned.To evaluate the differences in the mode of transferring labelled leukocytes, thedisappearance rate of labelled PMN in each recipient was estimated as the110relationship between the log of the fraction of PMNBrdU and time. The family of lineswithin each group were then compared using the restricted maximum likelihood(REML) method described by Feldman (Feldman, 1988). The estimates were thencompared using a chi-square statistic and differences in slopes, intercepts and lineswere considered to be significant when a probability than 0.05 was found. Using theestimated slope, the half-life was calculated for each group (Sachs, 1982).111RESULTSLabelling of PMN with BrdU:Figure 18 shows that a daily intravenous dose of 25 mg/kg of BrdU produced a rapidincreased in the percentage of BrdU labelled PMN. The immunocytochemicallabelling of PMN was variable and all cells with visible stain were deemed positive(figure 19a). The increased in BrdU labelled PMN occurred in a linear fashion fromday 1 to day 5 and gradually levelled out by day 7 when 80±2.3% of the PMN werelabelled. Immunoblot analysis of the DNA extracted from blood leukocyte samplesshowed that the amount of BrdU labelled DNA increased in a similar fashion (datanot shown).Washout of BrdU labelled PMN in recipient bloodTable VI shows that the number of PMN transferred as either LRP or as purifiedPMN was approximately one-half of the number of PMN in the same volume ofwhole blood. After the infusion of donor whole blood into recipients, the number ofBrdU positive PMN in the circulation, rose in an irregular fashion to a peak at 60minutes and then decreased rapidly over the next 24 hour period (figure 20a). WhenLRP (figure 20b) or purified PMN (figure 20c) was infused, the number of BrdUlabelled PMN in the recipient’s blood was lower at all the time points up to 60minutes.Calculation of the half-life of BrdU labelled PMNThe time required to achieve the maximal number of BrdU labelled PMN in thecirculating blood of the recipients who received whole blood was 60 minutes (figure20a). This time, was applied to the rate of decay equation to calculate the half-II2100Tz60II20T0 J0 1 2 4 5 6 7Time (days)Figure 18: Time course of BrdU incorporation into PMN. Blood samples were drawndaily from donor rabbits infused with 25 mg/kg/day of BrdU for 7 days. The fractionof BrdU positive PMN on cytospins prepared from LRP in donor blood is plottedagainst the days elapsed after the start of BrdU treatment. Day 0 represents thetime before the first injection. Each point represents the mean of three experiments±SE.“3Table VTCOMPARISON OF THE NUMBER OF WBC, PMN AND BrdU LABELLED PMNRECOVERED IN THREE DIFFERENT PREPARATIONS OF DONOR BLOODType preparation Number of leukocytes (x106)*WBC PMN BrdU labelled PMNWhole blood(8)t 257.7±8.8 116.3±8.9 85.9±22.1LRP (3) 132.3±1.8 52.5±3.8 41.6±3.9Purified PMN (6)- 60.0±5.0 46.2±3.1*lkt obtained from 25m1 donor blood.tnumber of samples analyzed.*average±SE.1/4Figure 19: Immunohistochemical detection of BrdU labelled cells. Panel (a)demonstrates a cytospin prepared from LRP of the donor blood after 7 days of BrdU25gfkg/day treatment. BrdU labelled PMN in the spleen (panel b) 24 hours afterinfusion of labelled PMN into recipient, the control lung (panel c) and Streptococcuspneumoniae infected lung (panel d) of recipients. BrdU labelled cells were transfusedas whole blood 3 hours after the instillation of either Streptococcus pneumoniae ora vehicle into the lung with colloidal carbon as marker. Bar =1Om.I.:,.100a II80 J0360Cz 40200 I0 120 2,40 360 1440 1450Time (mm)100b0380 J0360Cz L=; ‘i.3 200 120 240 360 1440 1450Time (miii)100C0?800?60bOC4020.1 00 I0 120 240 360 1440 1450Time (miii)Figure 20: BrdU labelled PMN in recipient’s circulation as a fraction of the numberof BrdU labelled PMN infused after the infusion of whole blood (19a, n=8), LRP (19b,open circles n=3) or purified PMN (19c, op.en circles n=3). Each data pointrepresents the mean with the SE. BrdU-PMN(% of injected) represents the numberof BrdU labelled PMN in the circulation as a fraction of the total number of BrdUlabelled PMN infused (see text).116life of PMN in the circulation:N = NTe -bwhere k represents the positive rate of loss of BrdU labelled PMN (slope),t the time after e = 2.7 1828, N the number of BrdU labelled PMN in thecirculation at time t and NT,,,X the number of BrdU labelled PMN in the circulationat time Tm.Since the half-life can be estimated as the time at which N = NTX /2, the rate-decayequation for the half-life, t112, becomes t112 = ln 2/k. The constant, k, was calculatedusing the restricted maximum likelihood method described by Feldman (1988). Theconfidence interval (Sachs, 1982) was obtained by deriving the lower and upperbounds of the 95 % confidence interval for the slope, k. Using this method ofcalculation, the half-life of the BrdU labelled PMN in the circulation of recipientrabbits using the whole blood infusion method was found to be 270 minutes with a95 % confidence interval of 248 and 296 minutes (table VII).Distribution of BrdU labelled PMN in recipient animalsTwenty-four hours after the recipients received the BrdU labelled cells, 55±11 % and58±9% of the original labelled PMN infused as either whole blood or LRP,respectively, were found in the spleen (Table VII, figure 19b). After the infusion ofpurified PMN, only 13±10% of the injected cells were recovered in the spleen. Veryfew BrdU labelled PMN were observed in sections of the liver, lung and bone117Table VIIHALF-LIVES OF BRDU LABELLED PMN INFUSED AS EITHER WHOLE BLOOD,LEUKOCYTE RICH PLASMA OR PURIFIED PMNType preparation slope halflife* 95% CIWhole Blood -0.0011132 270.4 248-296LRP -0.0011841 254.2 184-408Purified PMN -0.0024649 122.1 90-186* Half-life and confidence intervals (CI) are given in minutescalculated from the equation Nt=Naxed (see text)118marrow. In the lung and bone marrow the occasional labelled PMN was seen inmicrovessels. The point counting technique based on a grid density of 400 points was,however, not sensitive enough to quantitate the number of labelled PMN in theseorgans. The majority of BrdU labelled PMN seen in the liver were extravascular. Theanalysis of DNA extracted from these organs (figure 21) after infusion of whole bloodor LRP showed that the amount of BrdU labelled DNA in the spleen wassignificantly higher than that found elsewhere. In addition, Southern analysis of thelabelled DNA found in the spleen formed a ladder pattern typical of DNA from cellsundergoing programmed cell death or apoptosis (figure 22). This ladder pattern wasnot present in the DNA prior to infusion nor was it detected on the ethidium bromidestained gels of the total DNA extracted from the recipient spleens. BrdU labelledDNA was also found in the liver, bone marrow, gut and lung after whole blood orLRP infusion (figure 20). When adequate amounts of labelled DNA were present, aswas the case for the lung, the same ladder pattern found in the spleen was present(figure 22). After the infusion of purified PMN, BrdU was not detectable inequivalent amounts of DNA from these same organs, including the spleen.Activation parameters of BrdU labelled PMNL-selectin expression on BrdU labelled (82±8.5% labelled) and control PMN wassimilar as measured with indirect immunofluorescence using flow cytometry (MFI11.2 ± 2.8 versus 10.8 ± 3.1, BrdU labelled versus control PMN, n = 3). Similarly, CD 18expression was not different in BrdU labelled PMN. The ability of BrdU labelledPMN to generate superoxide with FMLP stimulation was the same as control PMN(7.1±3 versus 6.8±2.7nmolO 2/30 mm, BrdU versus controls, n=3). Likewise, withPMA stimulation, BrdU labelled and control PMN superoxide production wassiq.——4-4C).—0 ILiver Spleen Gut LungBone marrowOrganFigure 21: Semi-quantitation of BrdU labelled DNA in recipient organs 24 h afterthe transfusion of whole blood (solid bars, n=5) and LRP (hatched bars, n=3). TheSouthern blots were analyzed by densitometry and bars represent the averageintensity of immunological staining of DNA ± SE. Results were corrected for thenumber of BrdU labelled PMN infused.I2O4040 4 ,Figure 22: Immunological detection of BrdU labelled in DNA extracted from organsof one of the recipient rabbits after the transfusion of whole blood from a donorrabbit. A Southern blot of 20 g of DNA from each organ was stained for BrdU aswas 1Oig of DNA from donor peripheral blood used prior to transfer. Note the ladderpattern in the lung and the spleen lanes, indicating PMN apoptosis in these organs.Lanes are as indicated.121similar. These findings indicate that the labelling procedure did not activate orprime the labelled PMN. White cell, PMN and platelet counts did not change over the24 hour study period with any of the transfusion modes, indicating that thetransfusion per se did not affect the behavior of infused PMN.Migration of BrdU labelled PMN into inflammatory fociFour hours after the instillation of bacteria into the left lower lobe to inducepneumonia, there were more BrdU labelled PMN in the pneumonic area and BrdUlabelled PMN had migrated into the alveolar spaces (Figure 19c and 19d). The tissuesurrounding the subcutaneous sponge saturated with S. pneumoniae also containedmore intravascular and extravascular BrdU labelled PMN than the tissuesurrounding the control sponge.122DISCUSSIONLabelling PMN with radioisotopes ex vivo to study their turnover rates and behaviorin vivo has been well described (Denekamp and Kaliman, 1973; Doerschuk et aL,1987a; Muir et aL, 1984; Segal et at., 1976). These procedures have numerousdisadvantages and pitfalls, with one of the major concerns being the activation ofPMN during cell purification and labelling procedures (Haslett et aL, 1985; Hamiltonand Dobbin, 1983; McAfee et aL, 1984). Whether isotope labelled PMN represent thein vivo behavior of circulating PMN has been questioned. In the present study wehave demonstrated that BrdU can be used to label leukocytes in vivo and that theselabelled leukocytes can be successfully tranferred to recipient animals with minimummanipulation.In a previous study de Fazio et al (1987a) measured serum levels of BrdU in a mouseexposed to a dose of BrdU similar to what we have used. Steady state levels werewell in excess of circulating thymidine levels and no bone marrow toxicity could bedetected with their dosing regime over 7 days. They also demonstrated that thepercentage of cells labelled after a 30 mm pulse of BrdU was similar to valuesobtained for the S-phase percentage of normal mouse bone marrow as determined byflow cytometry, indicating satisfactory labelling efficiency with this dose of BrdU.Following 7 days of intravenous administration of BrdU, approximately 80 % of thecirculating PMN were labelled with BrdU. The intermittent daily dose of BrdU weused in our model may explain the ±20% BrdU negative PMN in the circulation aftera 7 day labelling period.123The first BrdU labelled PMN were detected in the circulation of the donors after 24hours with a small increase after 48 hours and a sharp rise to 28 ± 10% positive PMNwithin 72 hours (figure 17). This represents the minimum time from DNA synthesisin the last myelocyte generation cycle until the mature PMN are released in thecirculation. Using 3HTCIR the minimum transit time is 96-144 hours in humans(Fliender et at., 1964) and 48-72 hours in dogs (Maloney and Patt, 1968). Our studysuggests that the myelocyte blood transit time in rabbits is between 24-48 hours. Inthis study using a whole blood transfusion method the calculated half-life of the BrdUlabelled PMN in the circulation was 270 minutes or 4.5 hours (95 % CI 248-296minutes). This half-life is shorter than the 6.5 hours calculated in a previous studywhen PMN in rabbits were labelled with 51Cr (Doerschuk et at., 1987a) although inthat study purified PMN were used and the half-life was calculated from a singleexperiment. It is also shorter (between 6 and 10 hours) if compared to studies whereisotopes such as DP32P, ‘1In, [3HITdR and 51Cr were used in humans (Athens et at.,1961; Filender et at., 1964; Segal et al., 1976).Transfusion of whole blood was the most convenient and effective way of transferringthese labelled PMN to recipient animals. Using the same volume of blood, PMNpurification resulted in a calculated loss of approximately 50 % of the PMN due to thepurification procedure. Accelerated removal of these PMN from the circulation earlyfollowing infusion suggests they underwent changes during the purificationprocedure. Similar results were obtained when leukocyte rich plasma was infused.The most likely explanation for these observations is that the processing of PMNresults in sequestration of these cells and faster removal from the circulation pool124than PMN in whole blood (Haslett et al., 1985). Transfusing whole blood from thedonor to the recipient as described in this study avoids most of the problems that areencountered when labelling cells in vitro with isotopes.The in vivo labelling of PMN with BrdU did not activate the PMN, as wasdetermined by changes in the expression of two activation sensitive surface celladhesion molecules, L-selectin and CD 18, or prime the PMN, as was determined bytheir ability to produce superoxide. The functional capabilities of BrdU labelled PMNtransfused into recipients were further evaluated by their ability to migrate into fociof inflammation. We have demonstrated that BrdU labelled PMN accumulated in S.pnewnoniae generated inflammatory foci and migrated out of the microvessels of thelung into the alveolar space and out of systemic microvessels into infectedsubcutaneous tissue. This finding qualitatively assesses the ability of the BrdUlabelled PMN to migrate across the endothelium into the interstitium and/or alveolarspace in the lung. This preliminary evaluation of the functional capabilities of BrdUlabelled PMN suggests that the labelled PMN appeared to behave in a fashionsimilar to the unlabelled cells. The linearity of recruitment of BrdU labelled PMNin foci of inflammation should, however, be addressed in a quantitative study witha variety of stimuli in a dose-related manner, which is outside the scope of this thesis.It has been suggested that effete PMN leave the circulation randomly rather thanaccording to their age, do not re-enter the circulation (Fliedner et al., 1964a) andmeet their fate in situ (Hurley, 1983). The normal site for PMN emigration from the125circulation and death is unclear (Bainton, 1988) but our data show that over half ofthe BrdU labelled PMN infused as whole blood or LRP were found in the spleen 24hours after infusion. This figure contrasts with the 13±10% of BrdU labelled PMNrecovered in the spleens of the recipients who received purified PMN. This low valuefor removal of purified PMN by the spleen is similar to data obtained from a studythat used 51Cr labelled PMN in rabbits (Doersehuk et aL, 1987a). We attribute thisdifference to PMN activation during the isolation procedure, which results in rapidremoval of labelled cells by several organs, especially the liver. Previous studies with51Cr and 111n labelled PMN showed a substantial removal in the liver (Doerschuk etaL, 1987a; Muir et al., 1984). We suspect that the dispersion of the BrdU labelledPMN in an organ the size of the liver (100 times larger than the spleen in rabbits)makes it difficult to detect this shift of sequestration of PMN from the spleen to theliver using our morphometric method. It is also possible that a fraction of the PMNremoved into the tissues during the time frame of our experiment have already beendegraded and processed. The fate of PMN after they migrate into the tissues isunknown but they probably live for only 1 to 2 days (Bainton, 1988).The immunodetection methods used in conjunction with the Southern analysis of theDNA in this study showed the typical fragmented ladder pattern indicative of BrdUlabelled WBC undergoing programmed cell death (apoptosis) in the spleen and lung.The finding that apoptosis of WBC occurs in the spleen is consistent with thedisappearance and death of PMN in that organ. It is unlikely that apoptosis of otherleukocytes contributes significantly to the apoptosis pattern observed in the spleen.Lymphocytes are known to have a long half-life and the fraction of other granulocytes126in donor blood (<5%) is small. The observation that apoptosis also occurred in thelung is consistent with reports (Hogg, 1987) showing that there is a population ofPMN which have very long transit times in the lung. As the lung contains a largeproportion of the PMN marginated pool, it is conceivable that some PMN remainthere long enough to undergo apoptosis.This technique of labelling donor leukocytes in vivo and transferring the labelledcells to recipients in whole blood, provides a novel way to study PMN behavior invivo that is fast and simple. This method was used in the subsequent study todetermine the kinetics of L-selectin on PMN during their time spent in thecirculation.127Changes in L-selectin expression on PMN during their lifespan in thecirculationThe following specific aims wifi be address in this section:1) To transfused BrdU labelled PMN to recipient animals and detennine theexpression of L-selectin on these labelled PMN over time in the circulation.2) To determine whether the lack of L-selectin expression on circulating PMN serveas a signal for removal of these cells from the intravascular pool of PMN.IntroductionStudies on the behavior of PMN in the systemic circulation have shown that themigration of PMN out of the vasculature is preceded by the rolling of PMN on theendothelium of post capillary venules (Ley et al., 1991b; Von Andrian et al., 1991).mediated by the selectins (Lasky, 1992a). The leukocyte associated selectin, Lselectin, has been shown in vivo to mediate rolling of rabbit, rat and human PMN inmesenteric venules (Von Andrian et al., 1991; Ley et al., 199 ib). This rollingphenomenon allows PMN to marginate in post-capillary or collecting venules, is morepronounce on inflamed venules which allows the PMN to slow down and stop undershear force (Lawrence, and Springer, 1991; Smith et al., 1991). Roffing of PMN is anessential feature of PMN function in the presence of shear force both in vivo and invitro and may be facilitated by the conspicuous spatial distribution of the L-selectinon the tips of microvillus projections on the PMN cell surface (Picker et al., 1991).Rolling along the endothelium is followed by either a return of the PMN to thecirculation or by firm adherence in preparation for migration out of the vascularspace. This latter event is mediated by the activation-induced increases in avidity of128the integrins for their immunoglobulin-like counter receptors, intercellular adhesionmolecule (ICAM-l and ICAM-2) and vascular cell adhesion molecule (VCAM-l) on theendothelial cell surface (Butcher, 1991; Larson and Springer, 1990; Zimmerman etal., 1992).The only well described mechanism of regulation of L-selectin on PMN is theenzymatic cleavage of the receptor through biochemical processes that are still notwell understood (Kishimoto et al., 1989; Tedder, 1991). Stimulation of the receptor-bearing cells with activation-inducing agents or phorbol esters results in a shortperiod of enhanced avidity of L-selectin for its ligand on endothelial cells followed byimmediate receptor shedding (Kishimoto et al., 1989; Porteu and Nathan, 1990;Spertim et al., 1991b). This shedding of L-selectin may be essential to allow PMN tomigrate into inflamed tissues. The hypothesis that receptor-ligand interaction,involved in the rolling of PMN, results in the removal of L-selectin from the surfaceof PMN is attractive for several reasons: 1) the shedding of L-selectin provides a rapidmeans for the de-adhesion of the PMN allowing it to return to the circulation whenthe signal for firm adhesion is insufficient; 2) L-selectin can be found in thesupematant of cultured lymphocytes (Lanier et al., 1989; Spertim et al., 1991b); 3)recirculating lymphocytes shed L-selectin as they leave the vascular space to hometo peripheral lymph nodes (Gallatin et al., 1983a); 4) and L-selectin can bedemonstrated free in the plasma (Schleiffenbaum et al., 1992), suggesting that Lselectin may be constitutively lost from circulating leukocytes. Little is known aboutthe fate of L-selectin on circulating PMN, which do not recirculate and lack theability to synthesize L-selectin after leaving the bone marrow (Spertini et al., 199 ib).129The variable expression of L-selectin on circulating PMN in contrast to the highexpression on PMN released from the bone marrow further suggests that PMN loseL-selectin in the intravascular space. Doershuck et al (1994) have demonstrated thatjust a small number of PMN delivered to an area of pneumonia eventually emigrateinto the airspace, suggesting that the majority of PMN that interact with activatedendothelium in an inflammatory focus do not emigrate but return to the circulation.We conducted this study to test the hypothesis that PMN continuously shed L-selectinduring their stay in the circulation, resulting in older PMN expressing lower levelsof L-selectin. We further questioned whether the lack of L-selectin on the surface ofPMN signals their removal from the circulation. The study used the technique oftracing donor PMN labelled with BrdU in recipient animals (as has been describedabove), and allowed us to measure the expression of L-selectin on labelled PMN inrelation to the time PMN spent in the circulating blood.130METHODS:Animals:Twenty female New Zealand white rabbits were used in this study. Five (3.8 ± 0.3 kg,mean ± SD) were used as donors and 15 (2.3 ±0.3 kg) as recipients. The study wasapproved by the Animal Experimentation Committee of the University of BritishColumbia.Cell preparation:BrdU labelling of donor PMN:The DNA of rabbit leukocytes was labelled with BrdU, a thymidine analogue, aspreviously described (Bicknell et aL, 1994). Briefly, 5 donor rabbits were given BrdU(Sigma Chemical Co., St. Louis, MO) at a dose of 25 mg/kg daily for 7 days. Bloodwas collected from the central ear artery for blood cell counts, differential white cellcounts and to prepare LRP which was cytospun onto 3-aminopropyl-tri-ethoxysilanecoated slides. The cytospin specimens were air dried and stained using the APAAPmethod (Cordell et al., 1984) to determine the fraction of BrdU labelled PMN in eachspecimen (Bicknell et al., 1994).PMN punfication:The PMN were purified from donor rabbit blood as has been previously described(Doerschuk et al., 198Th). Briefly, LRP obtained from each 75 ml of donor blood wascentrifuged and resuspended in 1 ml PMN buffer. Hypotomc lysis of the residual redblood cells in the LRP was achieved by dilution with 11 ml sterile water. After 18seconds, 11 ml 2X PBS (2X PBS is 27 mM Na2HPO4, 132 mM KH2P04,and 2.74 MNaC1) and 10 ml PMN buffer were added. PMN were separated from the131mononuclear cells by centrifugation in Histopaque (Sigma Chemical Co.) with adensity of 1.077 g/ml at 150 x g for 13 minutes. The isolated PMN were 96% to 98%pure with a viability of 96 % as assessed by trypan blue exclusion.Removal of L-selectin by chymot,ypsin treatment of PMNSelective removal of L-selectin from PMN by chymotrypsin allowed us to study thebehavior of a population of predominantly L-selectin negative PMN in the circulation.To remove L-selectin from the surface of PMN we used the method described byJutila et al (Jutila et a!., 1991) who demonstrated that chymotrypsin cleave L-selectinfrom leukocyte surfaces without activating the PMN, as measured by changes inCD1 lb/CD 18, PMN random adhesion to plastic, migration to a chemotactic stimulusand changes in cell morphology measured by flow cytometry. Donor blood (75 ml) wasobtained and PMN were purified as previously described. The purified PMN weredivided into two specimens and were incubated in either denatured chymotrypsin(control) or chymotrypsin type IV (Sigma, St. Louis, MO) (diluted lU/mi for 3 x 106cells/lU incubated for 10 minutes at 37°C). PMN infused in control rabbits wereincubated with heat denatured chymotrypsin. In preliminary experiments the abilityof 5, 15, 30 and 60 mm heat denatured chymotrypsin to cleave a substrate N-BenzoylL-tyrosine ethyl ester was tested using a method described by Hummel et al (1959).The ability of chymotrypsin to cleaves the substrate was determinedspectrophotometrically with the use of a Perkin Elmer Lamda 2 UV/VISSpectrometer (Perkin Elmer & Co, Uberlinger, Germany). With 30 mm of heatdenatured chymotrypsin the ability of the enzyme to cleave the substrate wasabolished and this denaturing time was used in all subsequent experiments. The132efficiency of L-selectin removal by chymotrypsin from the PMN surface was evaluatedby two methods: a) immunocytochemically by using the APAAP method to stain forthe presence of L-selectin on isolated PMN cytospun on coated slides before and afterthe chymotrypsin treatment and b) immunofluorescence staining and flow cytometricanalysis of PMN before and after the chymotrypsin treatment (see below).In vitro effect of chymotrypsin on PMNThe effect of the chymotrypsin treatment on PMN activation parameters was testedto verify that the chymotrypsin treatment did not influence the behavior of PMN inthe circulation. Isolated PMN were divided into 3 samples, one was treated withchymotrypsin, one with denatured chymotrypsin and the remaining one incubatedwith buffer only before analysis.a) The expression of CD18 (MoAb 60.3, kind donation by Dr. J.M. Harlan) and Lselectin (MoAb DREG 200, kind donation by Dr. E.C. Butcher) on PMN weredetermined with immunofluorescent flow cytometric analysis.b) The viability of isolated PMN treated with chymotrypsin was also tested by thetrypan blue exclusion method which found no change from values before thechymotrypsin treatment.Experimental protocols:The leukocytes of donor rabbits were labelled in vivo with BrdU and these leukocyteswere transfused as either whole blood or purified PMN to serum compatiblerecipients. These BrdU labelled PMN were then followed in the circulation ofrecipients over a 24 hour period.133a) Twenty-five ml of donor whole blood (ACD as anticoagulant) or purified PMNobtained from 25m1 of whole blood (n=5) was transfused to serum compatiblerecipients. The disappearance of BrdU labelled PMN from the circulation wasmonitored over a 24 hour period by obtaining blood specimens from the central earartery at baseline (before infusion), at 2.5, 5, 7.5, 10, 15, 20, 30, 45, 60 mm and thenhourly until 6 hours after the infusion. A late specimen was collected after 24 hours.The white cell counts were determined on a Coulter Counter (Model SS8O, CoulterElectronics, Florida) and differential white cell counts on Wright’s stained bloodsmears. Leukocyte rich plasma was prepared from each specimen and cytospins weremade as described earlier.b) To determine the effect of removing L-selectin from the PMN surface on theirintravascular behavior, purified PMN from 75m1 of donor blood were diluted in 20m1PMN-buffer. The specimens were divided into two equal aliqouts which were thenincubated with either denatured chymotrypsin (control) or chymotrypsin type IV asdescribed earlier. The PMN treated in this way were then infused into the recipientsand the disappearance of BrdU labelled PMN from the circulation was monitoredover a 24 hour period by obtaining blood specimens from the central ear artery atbaseline, immediately after infusion of the PMN, 5, 10, 30, and 60 mm, and 3, 6 and24 hours later. White cell counts were obtained using a Coulter Counter (Model SS8O,Coulter Electronics, Florida) and differential white cell counts using Wright’s stainedblood smears. Leukocyte rich plasma was prepared from each specimen and cytospinswere made as described earlier.134Evaluating BrdU labelled PMN in recipientsDouble Immunoenzymatic staining of leukocytesLeukocytes on cytospins prepared from peripheral blood were stained for the presenceof surface L-selectin and nuclear BrdU using a double alkaline phosphatasetechnique. The APAAP technique (Cordefi et al., 1984) was used for both antigens.Surface L-selectin was first labelled followed by nuclear BrdU labelling. Briefly,slides were fixed in acetone for 10 mm, incubated with 5 % rabbit serum for 15minutes before the application of the monoclonal antibody against L-selectin, DREG-200 (5g/ml) for 60 mm in a humidity chamber at room temperature. Non-immunemouse IgG (5g/ml) and the omitting of the primary antibody was used as negativecontrols. As a linking antibody, a 1/20 dilution of rabbit anti-mouse IgG (Dako Z259)was applied for 45 minutes, followed by the anti-mouse alkaline phosphataseconjugated complex (Dako D651) in a 1/50 dilution for 45 minutes. All antibodieswere prepared in 50mM TrisCi, 150 mM NaCl, pH7.6 (TBS) with 1 % BSA and slideswere washed in TBS twice for 10 mm between each antibody application. Thealkaline phosphatase was developed by using a commercially available kit,HistoMark Red® (Kirkegaard and Perry, Gaithersburg, Maryland) for 10 minutes inthe dark. After a 10 minutes wash, slides were fixed 1 % paraformaldehyde for 10minutes. Cell membranes were further permeablized by methanol for 10 mm afterwhich time DNA was denatured by incubating slides in 2N HCL at 37°C for 60 mm.The 2N HCI was neutralized by washing the slides 3 times with 0. 1M borate buffer(BDH), pH 8.5. This was followed by the second APAAP procedure where mouse antiBrdU (Boeringher-Mannheim, Mannheim, Germany) 0. 1g/ml was used as theprimary antibody. With this procedure all slides were washed between antibody135applications with 0.1 % Tween 20 in TBS (pH 7.6) for 10 minutes. The alkalinephosphatase was developed with HistoMark Blue for 10 minutes in the dark. Slideswere washed for 30 minutes in distilled water, mounted in an aqueous medium(Gelvatol®) and analyzed on a Zeiss Universal Research light microscope (Model 1W,West Germany). The influence of the double labelling procedure on the presence ofsurface L-selectin and nuclear BrdU expression was evaluated by comparing thenumber of positive PMN for each antigen with paired slides stained for a singleantigen using the APAAP method as described above.Evaluating BrdU labelled PMNThe slides were coded and evaluated without knowledge of their origin. BrdU labelledPMN were evaluated on computer generated randomly selecting fields counting 100PMN per slide. BrdU labelled PMN were identified by a deep blue staining of the cellnucleus (figure 23). Cells were categorized as either BrdU positive (blue nucleus),BrdU and L-selectin positive (double labelled), L-selectin positive (red cell surface),or negative for both labels. If less than 10 % of the PMN were BrdU labelled, 200PMN were counted, if less than 5 %, 500 PMN were counted, and if less than 1 %,1000 PMN were counted. The total number BrdU labelled PMN evaluated in eachslide averaged 102± 19 with a range of 50 to 156. The number of BrdU labelled PMNpresent in the circulation of the recipients at each time point was expressed as afraction of the total number of labelled PMN infused corrected for the calculatedblood volume (American Physiology Societ, 1965) in the following manner:136Fraction PMNBrdU= PMNCWC X BV x %PMNe:emCirc BrdUPMNedwhere the fraction PMN represents the number of BrdU labelled PMN in thecirculation as a fraction of the total number of BrdU labelled PMN infused, PMN CfrCthe calculated number of PMN (1 x 106) in the circulation (total white cell count timesthe fraction of leukocytes that are PMN), BV the calculated blood volume, %PMNTthe fraction of BrdU labelled PMN in a cytospin of peripheral blood in the recipient,and PMN the number(1 x 106) of BrdU labelled PMN infused (PMN count/mi xml of fluid infused x %BrdU labelled PMN).The BrdIJ labelled PMN were further stratified as either stained for BrdU alone ordouble stained for BrdU and L-selectin, and these fractions were calculated separatelyusing the equation above.Calculation of the half-life of BrdU labelled PMNThe time required to achieve the maximal number of BrdU labelled PMN in thecirculating blood of the recipients who received purified PMN, Tmax, was applied tothe rate of decay equation to calculate the half-life of PMN in the circulation asdescribed in the previous section.‘4 .-.a‘37Figure 23: Immunocytochemical detection of BrdU labelled cells. Panel (a)demonstrates a cytospin prepared from LRP of the donor blood after 7 days of BrdU25gfkg/day treatments. Cells were stained for the presence of BrdU using theAPAAP method, with BrdU labelled cells staining red (see text). Panel (b)demonstrates double immunolabelling of PMN on cytospins for both nuclear BrdU(blue) and surface L-selectin (red) using a double alkaline phosphatase technique (seetext). Bar=lOMm.0IIb4138Statistical analysis:Statistical analysis was performed using SYSTAT® Version 5.1 software (Systat,Inc., Evanston, IL) (Wilkinson, 1990).A one-way analysis of variance was used to evaluate the changes in L-selectinexpression on PMN in the circulation over time, and a two-way analysis of variancewas used to compare the transfusion of whole blood with isolated PMN over time.To evaluate the differences in the behavior of chymotrypsin-treated or control BrdUlabelled PMN the disappearance rate of labelled PMN in each recipient wasestimated as the relationship between the log of the fraction of pBrdU and time.The family of lines for the individual animals within each group were then comparedusing the restricted maximum likelihood (REML) method (Feldman, 1988). Theestimates were then compared using a chi-square statistic and differences in slopes,intercepts and lines were considered significant when the p-value was less than 0.05.The estimated slope was then used to calculate the half-life for each group (Bryan etaL, 1990).139RESULTSIn vivo labeffing of PMN with BrdUDaily intravenous injections of 25 mg/kg of BrdU produced a rapid increase in thepercentage of BrdU labelled of the PMN in donor rabbits with 82±4% PMN labelledby 7 days. AU cells with visible nuclear BrdU stain were deemed positive (figure 23a).In double labelled slides, the surface L-selectin was stained red and the nuclear BrdUblue with overlapping areas stained purple (figure 23b). Expression of both L-selectinand BrdU was similar in paired single and double stained slides.L-selectin changes on PMN during their intravascular lifespanThe fraction of L-selectin negative BrdU labelled PMN increased with time spent inthe circulation whether or not the PMN were infused as whole blood (p <0.001) or aspurified PMN (p <0.0001) (figure 24). The fraction of BrdU labelled L-selectinnegative PMN infused was higher for the purified PMN (14.2±1.9%) than for thewhole blood (7.5±2%), resulting in a higher fraction of L-selectin negative BrdUlabelled PMN in the recipient circulation directly after purified PMN were infused(23±5.2%) than after the infusion of whole blood (11±2.6%). This difference wasattributed to a small loss of L-selectin from PMN during the purification process,which was confirmed with flow cytometric analysis of these populations of PMN(figure 25a and b). In both transfusion modes, a rapid increase in L-selectin negativePMN occurs within the first 30 mm after infusion followed by a more gradualincrease over the rest of the study period. After 24 hours, with the infusion of eitherwhole blood or purified PMN, nearly all the PMN were negative for L-selectin (figure24)./4’Oz100-I.80-CT60- ,o-.. - -. - -/ I0 80 160 240 320 400400 1440Time (mm)• Whole Blood - -0 - Purified PMNFigure 24: L-selectin changes on PMN during time spent in the circulation whenBrdU labelled PMN were infused as whole blood (closed circles, n=5) and purifiedPMN (open circles, n=5). Values are the mean±SE. PMN on cytospins made ofleukocyte rich plasma were double immunolabelled for BrdU and L-selectin (see text).The fraction of L-selectin negative BrdU labelled PMN (y-axis) increased significantlyover time in both groups (p <0.001) with nearly all the BrdU labelled PMN beingnegative for L-selectin after 24 hours.4-’D0C.)a)0Mean Fluorescence Intensity14!Figure 25: The effect of chymotrypsin (lU/mi for 3x106 PMN incubated for 15 miii)on the expression of L-selectin (a, b and c) and CD18 Cd, e and f) as measured withflow cytometry. Panels a and d represent baseline expression of PMN L-selectin andCD18 respectively determined on whole blood. Panels b and e represent similarexpression after PMN purification (note the small loss in L-selectin with PMNpurification). Panels c and f represent PMN L-selectin and CD18 expression afterchymotrypsin treatment respectively. Note the significant loss of L-selectin from thePMN with chymotrypsin treatment but no changes in the expression of CD18. Inpanels the events left of cursor 1 represent non-specific labelling. The x-axisrepresents events and the y-axis the mean fluorescence intensity plotted on a log-a b c__________I1 1d e fI I_1 1.•1scale.142In vitro effect of chymotrypsm on PMNCleavage of L-selectin from PMN by chymotrypsinThere was no difference between the expression of L-selectin on PMN incubated indenatured chymotrypsin or buffer alone, therefore all subsequent results are fromPMN incubated in denatured chymotrypsin before analysis and infusion. Figure 25band 25c demonstrated that 78±8.2% of the L-selectin present on PMN was lostfollowing chymotrypsin treatment. This loss was confirmed withimmunocytochemistry by grading the expression of L-selectin on PMN present incytospins prepared from leukocyte rich plasma, where PMN highly positive for Lselectin decreased from 63 ± 6.8 % to 4.6 ± 3 % with chymotrypsin treatment.Effect of chymotrypsin on PMN activation parametersThe viability of PMN, accessed by the trypan blue exclusion, was the same before andafter chymotrypsin treatment and the treatment did not change the expression ofCD18 (figure 25d and 25e) as measure by flow cytometry.The clearance of chymotrypsin treated PMN from the circulationThe fraction of BrdU labelled PMN present in the circulation of recipients directlyafter infusion was the same in both the control (48±12.6%) and the chymotrypsintreated (46±9.6%) groups. Although the number of chymotrypsin treated PMNdecreased in the first hour following infusion, they returned to the circulation by 3hours and then disappeared from the circulation at the same rate as in the controlgroup (figure 26). This similarity was reflected in the overall half-lives (262 versus296 minutes) of both groups (table Vifi). In both groups L-selectin positive PMN were100C)0 80.4-060z0 80 160 240 320 4001400 1500Time (mm)0 Control—• ChymotrypsinFigure 26: BrdU labelled PMN in the circulation of recipients after the infusion ofpurified PMN treated with either denatured chymotrypsin (open circles, n=5) orchymotrypsin (closed circles, n=5). PMN were incubated withchymotrypsinldenatured chymotrypsin (1U/ml for 3 x 106 PMN) for 15 mm. Valuesare mean±SE of the number of BrdU labelled PMN in the circulation as a fractionof the number of BrdU labelled PMN infused.100 144aU)0U) 80060za40 ToK--- I0 80 160 240 320 4001400100 b04-0o 80t060za• 40TS...,20.--- T0 7—0 80 160 240 320 4001400Time (mm)0 L-seIectin+ —-•--— L-seiectinFigure 27: BrdU labelled PMN in the circulation of recipients after the infusion ofpurified PMN treated with either denatured chymotrypsin (a, n=5) or chymotrypsin(b, n=5). PMN were incubated with chymotrypsinldenatured chymotrypsin (1U/ml for3x106PMN) for 15 mm. Values are mean±SE of the number of BrdU labelled PMNin the circulation expressed as fraction of the number of BrdU labelled PMN infused.The closed circles represent the washout of L-selectin negative PMN and the opencircles L-selectin positive PMEN. In both groups L-selectin positive PMN clear fromthe circulation faster than L-selectin negative PMN. With chymotrypsin treated PMN(b), L-selectin positive PMN cleared from the circulation significantly faster than Lselectin negative PMN (p < 0.05).145cleared from the circulation faster than L-selectin negative PMN (figure 27a and27b). In the chymotrypsin group, L-selectin positive PMN were cleared from thecirculation faster than L-selectin negative PMN. However, the half-lives of L-selectinnegative PMN in both group did not differ from that of the half-life of the wholepopulation of PMN (table VIII).‘4’TABLE VifiHalf-life of PMN treated with and without ChymotrypsinCell populations T112 Confidence intervalsUpper(min) Lower(min)Control PMN’;All 262 458 184L-selectin+ 178 311 134L-selectin- 219 320 135Chymotrypsin treated PMN;All 296 570 199L-selectin+ 133 238 118L-selectin- 302 722 180*#Control PMN were incubated with denatured chymotrypsin*Half..life of the L-selectin negative PMN was significantly longer than the L-selectinpositive PMN in the group that was infused with chymotrypsin treated PMN.147DISCUSSIONThe results presented here show that the fraction of L-selectin negative PMNincreases during their stay in the circulation. This may be either from L-selectinpositive PMN losing their L-selectin in the circulation and changing to negative cellsor from the preferential removal of L-selectin positive PMN from the circulation. Ourdata favour the hypothesis that L-selectin is progressively lost from the surface ofPMN as they marginate and demarginate in the circulation, resulting in older PMNexpressing lower levels of L-selectin. We also demonstrated that cleaving L-selectinfrom the surface of PMN with chymotrypsin did not result in the accelerated removalof these PMN from the circulation.The adhesive interaction between leukocytes and endothelial cells involves multiplesequential steps (Butcher, 1991; Lawrence and Springer, 1991). The initial adhesivestep is leukocyte rolling which is mediated by members of the selectin family ofadhesion molecules (Abbassi et al., 1993; Butcher, 1991; Lawrence and Springer,1991) that recognize fucosylated carbohydrate ligands, especially structurescontaining sialylLewisx (sLex) (Lasky, 1992a; Springer, 1990b). L-selectin isconstitutively expressed on nearly all circulating PMN (Griffin et al., 1990; Tedderet aL, 1990a) that interact with endothelial cell surface ligands, resulting in therolling and margination of PMN in post-capillary venules. This rolling phenomenonhas been shown to occur or normal unactivated endothelium (Ley et al., 1993) but isenhanced on activated endothelium with L-selectin presenting sLex to the inducibleendothelium-associated selectins, P- and E-selectin (Picker et al., 1991; Spertini et al.,1991a). The fraction of leukocyte rolling increases from under 20% on unactivated148endothelium to over 30 % on cytokine activated endothelium with a marked decreasein rolling velocity (Ley et al., 1993; Ley et al., 1991b). Rolling cells become activatedwhen they encounter appropriate activating or chemotactic stimuli generated on theendothelial surface or in the extravascular compartment. PMN activation results inboth functional activation and up-regulation of the 32-integrins (CD 11/CD 18) andallows firm adhesion between PMN and the endothelium mediated via theCD18/ICAM-1 system (Arfors et al., 1987; Lawrence and Springer, 1991). PMNactivation has also been shown to result in the shedding of L-selectin as a deadhesiveevent. This chain of events has been shown in vivo to be essential for PMN toemigrate into the extravascular space toward an inflammatory focus (Arfors et al.,1987; Von Andnan et al., 1992).The principle finding in this study is that PMN L-selectin expression decreases withtime spent in the circulation, which may be either from L-selectin positive PMNlosing their L-selectin in the circulation and changing to negative cells or from thepreferential removal of L-selectin positive PMN from the circulation. Our data favorthe concept that PMN shed L-selectin during their intravascular life becauseshedding is the principal mechanism for PMN to regulate their L-selectin expression(Spertini et al., 1991b; Tedder, 1991). It is conceivable that PMN could shed their Lselectin when they encounter mildly activated vascular beds such as the gums, upperrespiratory tract and bladder where they are inadequately stimulated to support firmadherence and migration and that PMN released from these vascular beds couldexpress lower levels of L-selectin. Previous studies from our laboratory havedemonstrated intravascular activation of PMN with shedding of L-selectin and* 149without PMN migration in the lungs of rabbits who had been exposed to cigarettesmoke and complement fractions (Doerschuk and Allard, 1989; Kiute et aL, 1993).However, this is unlikely to be the major mechanism for the loss of L-selectinexpression on the PMN in our study because the size of these mildly activatedvascular beds is small in relation to the total microvascular surface available.Spontaneous shedding of L-selectin in lymphocytes has been demonstrated in cultureby Spertini et al (1991b), but whether PMN behave similarly is unknown.Furthermore, high levels of circulating free L-selectin (sL-selectin) in plasma havebeen demonstrated by Schleiffenbaum et al (1992), suggesting that leukocytescontinuously shed L-selectin while while they in the intravascular space. A moreattractive hypothesis is that ligand binding initiates receptor shedding whichprovides a rapid means for the regulation of leukocyte adhesion to endothelial cellswith the subsequent release of the cell allowing de-adhesion. This de-adhesion eventmay be necessary to allow the to PMN return to the circulation if the conditions forfirm adhesion and migration are inadequate or alternatively to migrate out of thevascular space at inflamed sites. The stage at which L-selectin is shed duringleukocyte-endothelial interaction has not been established. Hypothetically, the rollingof PMN on “normal” or activated endothelium, the exposure of PMN to endotheliumderived activating factor, or the migration process through the endothelium itselfmay all be associated with L-selectin shedding. Shedding may also occur at all abovementioned stages of leukocyte-endothelial interaction in a graded fashion. Our resultsshow that normal circulating PMN decrease their expression of L-selectin over timeand that PMN that remaining in the circulation for 24 hours are universally L150selectin negative. This finding suggests that the normal contact between PMN andthe endothelium may result in a loss of surface L-selectin.Technical factors could contributing to the decrease in PMN L-selectin expressionseen in our model specifically, activation of the PMN by either the labeffing,purification or transfer procedures. We have shown though that labelling PMN withBrdU did not activate or prime the cells (previous section). This method of labellingPMN allowed us to study the behavior of an activation sensitive surface molecule,such as L-selectin, over time in the circulation. It is thus unlikely that the labeffingprocedure per se result in PMN activation.The purification procedure of PMN resulted in a small loss of PMN L-selectin (figure25a and b), suggesting that the PMN were mildly activated during the purificationprocedure. However, fact that CD18 expression did not change that would argueagainst significant cell activation. This finding is in contrast to the response ofhuman PMN to purification (Kuijpers et al., 1991), where CD18 expression increaseswith PMN purification and L-selectin expression remains unchanged. Activated PMNhave been shown to behave differently in the circulation than normal unactivatedcells and are removed from the circulation faster (Haslett et al., 1985). However, inthis series of experiments the removal of purified PMN (262 mm 95 % CI 458-184) wassimilar to values obtained when PMN were infused as whole blood (270 mm 95 % CI248-296).It is unlikely that the transfusion per se influenced the PMN behavior as both whole151blood and the purified PMN were given to serum-compatible recipients and neitherproduced changes in total white cell, PMN, band cell and platelet counts or increasedCD18 expression on PMN over the 24 hour study period (data not shown). Weconclude that the labelling, purification and transfusing procedures did notsignificantly activated the PMN.To test the hypothesis that the lack of L-selectin on the surface of PMN is a signalfor their removal from the circulation, we selectively removed L-selectin from thesurface of PMN. We have shown that PMN L-selectin expression decreases over timein the circulation which implies that L-selectin negative PMN represent an olderpopulation of PMN. An older population of PMN should theoretically have a shorterhalf-life than newly released cells from the bone marrow. Removing L-selectin fromPMN with chymotrypsin (Jutila et al., 1991) greatly increased the population of Lselectin negative PMN we transfused (figure 25). After infusing these treated PMNinto recipients, an early temporary removal of these cells from the circulating poolwas seen, but they reappeared in the circulation between 1 and 3 hours. Additionally,similar half-lives were calculated for the control and the chymotrypsin treatedgroups. These findings suggest that the lack of L-selectin on PMN did not result intheir permanent removal from the intravascular compartment.The calculated half-lives of the L-selectin negative PMN in the control and thechymotrypsin treated groups were similar if compared to the whole population oflabelled PMN (table VIII). This longer than expected half-life of the L-selectinnegative PMN may represent a continuous shift of L-selectin positive to L-selectinnegative PMN in the circulation, which may also account for the shorter half-lives152of the L-selectin positive PMN in both the control and the chymotrypsin treatedgroups.The daily turnover rate of PMN in humans is 100 to 200 billion cells, with themajority of these PMN being removed by the spleen and liver (Andrewes, 19 lOb;Andrewes, 1910a; Bicknell et al., 1994). Our results suggest that a lack of L-selectinexpression is not the signal responsible for the removal of these older PMN from thecirculation because removing L-selectin from PMN with chymotrypsin, simulating“older” PMN, does not result in the accelerated removal of these cells from thecirculation.Our data show a smaller fraction of L-selectin positive than negative PMN in thefirst specimen of circulating blood taken after the infusion of labelled PMN (figure27a and b). This finding suggests that the L-selectin positive PMN are preferentiallyremoved from the circulating pooi of blood early after infusion and is consistent withprevious studies demonstrating preferential sequestration of L-selectin positive PMNin the lungs of rabbits exposed to zymosan activated plasma (Graham et aL, 1992;Quinlan et al., 1992). Our findings extend these observations by showing that Lselectin positive PMN preferentially marginate in normal unstimulated animals.Moreover, a rapid increase in L-selectin negative PMN was seen within the first 30mm after the infusion of labelled whole blood and purified PMN (figure 24), whichmay represent preferential margination of L-selectin positive PMN. Taken together,these findings suggest preferential margination of L-selectin positive PMN thatreturn to the circulation with less L-selectin resulting in an increasing pool of153circulating L-selectin negative PMN.Other studies from our laboratory have demonstrated that in normal circulatingPMN in humans (previous sections) and rabbits (Klute et aL, 1993) the expression ofL-selectin is variable with populations of PMN expressing high, intermediate and lowlevels of L-selectin, These PMN contrast with mature PMN in the bone marrowwhere the majority express high levels of L-selectin (Lund-Johansen and Terstappen,1993; previous section) resulting in an increase in L-selectin expression on circulatingPMN with the bone marrow release of PMN. It has been shown in vitro and in vivothat in the chain of events responsible for leukocyte-endothelial interaction, L-selectinis essential for PMN emigration to an area of inflammation (Ley et al., 1991b;Lund-Johansen, and Terstappen, 1993; Mulligan et a!., 1991; Mulligan et al., 1994;Tate and Repine, 1983a), implying that PMN recently released from the bone marroware equipped to be preferentially recruited to foci of inflammation whereas olderPMN with less L-selectin may be less able to be recruited to these sites. Reducedlevels of L-selectin on neonatal PMN (±50% of adult values) are related to theirreduced ability to adhere under flow conditions to activated endothelium (Andersonet aL, 1991). This neonatal PMN deficit in L-selectin further emphasizes theimportance of a critical receptor density for L-selectin-dependent function. Wespeculate that PMN with reduced or absent L-selectin represent a subpopulation ofolder circulating PMN with reduced functional capabilities, such as their ability tomarginate and emigrate to areas of inflammation.1546) SUMMARY AN]) FUTURE DIRECTIONSThe emigration of PMN to sites of inflammation requires intercellular adhesion.During recent years there has been an extraordinary increase in informationconcerning the mechanisms that mediate this leukocyte-endothelial cell interactionduring inflammation (Butcher, 1991; McEver, 1992 Springer, 1990b; Zimmerman etaL, 1992; Yong, and Khwaya, 1990). The cell adhesion molecules play a pivotal rolein this interaction during inflammation in the systemic and pulmonary microvessels.However, the role of these molecules in the trafficking of PMN from the bone marrowinto the blood and changes during their lifespan in the circulation are less clear. Inthis thesis we have explored the fate of the cell adhesion molecule, L-selectin, onPMN from the time of their release from the bone marrow into the bone marrowvenous sinusoids and the general circulation until their permanent removal from theintravascular pool (figure 28). We have demonstrated that the mature segmentedPMN in the bone marrow hematopoietic tissue express the highest levels of Lselectin. Some of this L-selectin is shed when these cells cross the bone marrow-blood-barrier. During a chemotactic stimulus of the bone marrow the barrier may becomemore permeable and this loss of L-selectin may be less marked. Under theseconditions the expression of L-selectin on circulating PMN nearly equals theexpression on bone marrow PMN, although the same level of expression is neverreached. This also suggests that the PMN expressing high levels of L-selectin in thecirculation at baseline are newly released PMN from the bone marrow. We havefurther demonstrated that PMN in the circulation progressively lose L-selectin thelonger they remain in the circulation. This finding may explain the variable 7’expression of L-selectin on circulating PMN at baseline, with newly released PML-selectin during bone marrow release of PMN’sBaselinePrecursorBone marrowMature PMN•zzBone Marrow sinusoi .. Circulating blood* edding• L-seleCtifl Tissue156expressing the highest and older PMN the lowest levels of L-selectin. This lack of Lselectin on older PMN in the circulation has been shown to be not a signal for theirremoval from the circulation.It is known that many chemotactic factors play a key role in leukocyte homeostasis,particularly in the setting of inflammation. These factors also stimulate the releaseof PMN from the bone marrow which functionally links the initial neutropenia,chemotaxis and eventual neutrophilia as a continuous dynamic process that is self-regulated at the cellular level. The precise mechanism(s) leading to neutrophilicleukocytosis at the cellular level remain undefined. Adhesion events between theegressing cells and the bone marrow microenvironment may play a key role in thisdynamic process. In leukemia for example, immature white cells are able totransverse the bone marrow-blood barrier and appear in the blood in contrast to theirnormal immature counterparts. Even when there is a relative increase in myeloblastand promyeloblasts as compared to differentiated granulocytes, as seen withagranulocytosis, there is no egress of immature forms into the blood. The exceptionoccurs when recombinant hematopoietic growth factors (GM-CSF) are given topatients in pharmacological doses. These factors also alter the adhesive properties ofleukocytes which may contribute to their ability to egress from the bone marrow.Similarly, adhesive properties of leukemic cells differ from normal immatureleukocytes that may explain their ability to egress from the bone marrow. Both GMCSF stimulated and acute and chronic myeloid leukemic cells express low levels ofL-selectin (Demetri and Antman, 1992; Spertini et al., 1991b). This lack of theadhesion molecule may be partially responsible for their relocation from the bone157marrow into the circulation. In aleukemic leukemia the blast cells have differentpotentials to gain access to the circulation and it is tempting to speculate that thesecells have similar adhesive qualities as “normal” myeloblasts. The development of aL-selectin knock-out animal may shed further light on the role of L-selectin in theegress of PMN from the bone marrow.Our finding that intravascular complement activation stimulates the bone marrowto release PMN expressing high levels of L-selectin may have clinical implications.Polymorphonuclear leukocytes have been implicated in the pathogenesis of severalimportant clinical conditions that including lung destruction in emphysema, adultrespiratory distress syndrome (ARDS), ischaemia-reperfusion injury in the gut andthe heart and multi-organ failure associated with septic shock (Hernandez et al.,1983; Tate and Repine, 1983a; Tennenberg et al., 1987; Senior et aL, 1977). Theincrease in circulating segmented and non-segmented PMN from the bone marrowstores has been attributed to a variety of stimuli (Jagels and Hugli, 1992; Marsh etal., 1967; Ulich et al., 1989). Chronic cigarette smoking for example, produces a 20%-25 % increase in peripheral blood leukocyte counts compared to non-smoking controlswhich may be related to excess marrow release (Corre et al., 1971; Hunninghake andCrystal, 1983; Janoff, 1983). Recent evidence suggests that PMN in the bone marrowpool are mobilized after intravascular complement activation in rabbits (Doerschukand English, 1991) and sheep (Rosolia et aL, 1992). When PMN are released from thebone marrow following complement activation (Doerschuk and English, 1991; Rosoliaet al., 1992), the PMN expressing high levels of L-selectin have been shown topreferentially sequester in the lungs (Quinlan et al., 1992; Graham et al., 1992). In158this study we have shown that newly released PMN express high levels of L-selectin,and it is tempting to speculate that these newly released PMN preferentiallymarginate and adhere to activated endothelium and emigrate into inflammatory foci.Therefore, they could play a critical role in the pathogenesis of PMN-induced organinjury such as ARDS and multi-organ failure, and in doing so are a prime target fortherapeutic intervention in conditions of PMN mediated tissue injury.Our finding that newly released PMN express high levels and older PMN lower levelsof L-selectin, creates the opportunity to selectively study the functional capabilitiesand trafficking of these different populations of circulating PMN. The concept thatthe circulating pool of PMN are a uniform population of cells and that these cells arerandomly removed from the vascular space can now be tested directly.1597) REFERENCESAbbassi, 0., Kishimoto, T.K., Mclntire, L.V., and Smith, C.W. (1993). 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