Impairment of HIV polymorphonuclear leukocyte transmigration across T84 cell monolayers: an alternative mechanisms for increased intestinal bacterial infections in AIDS?

ARTICLE

INTRODUCTION

Numerous PMN functions are impaired in all stages of HIV infection. Qualitative functional defects of HIV PMN have been observed in vitro, including defects in phagocytosis, superoxide production, as well as accelerated apoptosis [1-9]. Concerning the chemotaxis of HIV PMN, previous studies have generated conflicting results showing decreased, increased, or even normal function [4, 10]. Diarrhea is a common symptom in patients infected with HIV; however a substantial proportion of patients have no etiological explanation for their symptoms after diagnostic evaluation, reaching 50% in some series [11, 12]. In these patients, nonspecific inflammation may be seen on mucosal biopsy [13, 14]. On the other hand, the incidence of bacterial infections of the gastrointestinal tract such as salmonellosis, shigellosis or colibacillosis, is high in AIDS patients, even in the early stages of the disease [15-18]. Several authors have suggested that intestinal inflammation could be a direct consequence of HIV infection and previous studies have established that the gastrointestinal tract contains cellular reservoirs of HIV [19, 20]. The colorectal mucosal surface epithelium can be infected by the HIV which could induced some cellular modifications, increasing the susceptibility of intestinal bacterial infections [19, 20]. However, a diminished transepithelial migration of PMN could lead to decreased bacterial phagocytosis, promoting bacterial proliferation and epithelial invasion by the bacteria. The aim of this study was to compare, using an in vitro model [23], the migration of control PMN and HIV PMN across an intestinal epithelium barrier. We used the T84 cells which exhibit the functional and morphological characteristics of colonic cryptic cells [24]. Recruitment of PMN to sites of inflammation requires adhesive interactions with different surfaces (endothelium, extracellular matrix, epithelium). A major part of these interactions is mediated by binding of the conformationally activated ß2 integrin Mac-1 (CD11b/CD18) to specific ligands [25]. More specifically, transepithelial migration of PMN has been shown to be a CD11b/CD18-dependent event, as evidenced by the inhibitory effect of CD11b/CD18 antibodies on the transmigration process [26]. Recently, it has been demonstrated that the integrin-associated protein known as CD47 is involved in neutrophil transmigration across intestinal epithelium [27]. CD47 is also involved in the host defense against bacterial infection [28]. In this study we have investigated, via flow cytometry, the expression of CD11b, CD18 and CD47 on control PMN and HIV PMN, on non-transmigrated cells and cells obtained after transepithelial migration. Using electron microscopy we compared the expression of CD11b on control PMN and HIV PMN, with and without stimulation by the formyl-met-leu-phe peptide (f-MLP). Finally, as accelerated apoptosis may increase the risk of secondary infections, we investigated and compared apoptosis of transmigrated HIV PMN and control PMN.

SUBJECTS AND METHODS

Study population

Forty-five HIV-positive individuals aged 27-67 years (mean, 42 years; 25 men and 20 women) were included in the study. Risk factors for HIV infection included homosexual contact for 17 (37%), intravenous drug use for 18 (40%; 6 men and 12 women), heterosexual contact for 6 (13%; 3 men and 3 women), and blood transfusion for 3 (8%; 2 men and 1 women). Risk factors were unknown for one man (2%). Twenty-seven patients had asymptomatic HIV infection (Center for Disease Control and Prevention (CDC) class II) (Group 1) [29]. Eighteen patients had an AIDS-defining disease according to the CDC classification with a CD4⁺ cell count < 200 x 10⁶/l (Group 2) [29]. Twenty-six blood samples were obtained from patients who had received various medication as prophylaxis against opportunistic infections and/or treatment of HIV. None of the patients had neutropenia or concurrent infection during this study. Forty-five HIV-negative individuals, aged 25-35 years, served as a control population.

Tissue culture

T84 cells, (ATCC, passages 65-90), a human colonic carcinoma cell line, were grown and maintained as confluent monolayers on collagen-coated, permeable supports with detailed modifications [23]. T84 cells were grown as monolayers in a 1:1 mixture of Dulbecco-Vogt modified Eagle's media (DMEM) and Hanks F-12 medium supplemented with 15 mM N-2 hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) buffer (pH 7.5), 14 mM NaHCO₃, 40 mg of penicillin per ml, 90 mg of streptomycin per ml, 8 mg of ampicillin per ml and 5% newborn calf serum. Monolayers were grown on 0.33-cm² ring-supported polycarbonate filters (Costar, Cambridge, Mass.) and utilized 6 to 14 days after plating. Steady-state resistance was reached in 4 to 6 days, with variability largely related to cell passage number. Monolayers received one weekly feeding following initial plating. Confluent monolayers on permeable supports were constructed to permit a basolateral-to-apical migration of PMN ("inverted inserts") as previously described [23].

Preparation of neutrophils and neutrophil transmigration assay

Human neutrophils were isolated from whole blood using a gelatin-sedimentation technique [26]. Briefly, whole blood anticoagulated with citrate/dextrose was centrifuged at 300 x g for 20 min (20° C). The plasma and buffy coat were removed and the gelatin/cell mixture was incubated at 37° C for 30 min to remove contaminating red blood cells (RBC). Residual RBC were then lysed with isotonic ammonium choride. After washing in HBSS without Ca²⁺ or Mg²⁺, the cells were counted and resuspended at 5 x 10⁷ PMN/ml. PMN (95% pure) with 98% viability by trypan blue exclusion were used within 1 hour after isolation.

The physiologically (basolateral-to-apical) directed PMN transepithelial migration assay has been previously described [23]. Neutrophil transmigration experiments were performed at 37° C on 0.33-cm² inverts. Once isolated, the PMN were suspended in modified HBSS (without Ca²⁺ and Mg²⁺, with 10 mM HEPES (pH 7.4; Sigma Chemical Co) at a concentration of 5 x 10⁷/ml. We added 1 x 10⁶ PMN to the inverts. Transmigration of PMN was initiated by different concentrations f-MLP (10^-7, 10^-8, 10^­9 M) to the lower reservoir or 100 ng/ml IL-8 (Genentech) and incubating it for 15 min to allow a transepithelial chemotactic gradient to form prior to the addition of PMN. Transmigration of neutrophils was assayed by quantification of the azurophil granule marker myeloperoxidase (MPO) as described previously. Briefly, after transmigration, T84 monolayers were rapidly cooled to 4° C, washed with HBSS and solubilized in 1% Triton X-100-containing HBSS. The pH was adjusted to 4.2 with a 1:10 dilution of 1.0 M Na citrate, pH 4.2, and peroxidase activity was assayed by the addition of an equal volume of 1 mM 2,2'-azino-di-(3-ethyl) dithiazoline sulfonic acid and 10 mM H₂O₂ in 100 mM citrate, pH 4.2. To quantify neutrophils which transmigrated through the monolayer into the lower reservoir, 1% Triton X-100 was added directly to the reservoir and assayed as above.

Immunoelectron microscopic study

For immunoelectron microscopy, resting PMN pellets or f-MLP stimulated PMN pellets (30 min, 10^­7M f-MLP) were fixed in 3.7% paraformaldhehyde and embedded at low temperature into LR White resin (Hard LR White, London, UK). Ultrathin sections were put on 300 mesh nickel grids, washed with phosphate-buffered saline (PBS), then incubated for 60 minutes at room temperature with CD11b antibody (OKM1; ATCC, diluted: 1/100). After washing with PBS, the grids were incubated for 60 min with 10 nm colloidal gold-conjugated rabbit anti-mouse secondary antibody (TEBU, Paris, France). The grids were washed with PBS, then with distilled water and stained with uranyl acetate. Sections were examined with a JEOL 1200 EXII electron microscope. The total number of beads present per PMN were counted at the cell surface membrane in a random section for each conditions. These counts were performed on 30 PMN per condition.

Flow cytometric assay

Flow cytometric assay was performed before and after 2 hours transmigration at 37° C. Neutrophils that had transmigrated through the epithelial monolayer from 12 Costar plates (Low attachment, Cambridge, MA) were pooled for flow cytometric analysis as well as control PMN in HBSS with and without f-MLP (10^­7 M, 2 hours). Neutrophils in HBSS were fixed in 1% formalin for 30 min at room temperature. The cells were then washed once in HBSS and incubated with polyclonal goat Ig for 20 min. The neutrophils were washed again in HBSS and treated with either mAb OKM1 (anti-CD11b) (ATCC; diluted: 1/1000), mAb BRIC126 (anti-CD47)(International Blood Group reference Laboratory, Bristol, UK; diluted: 1/500) and an isotype-matched control, or HBSS for 20 min at room temperature and then washed twice. Cells were then exposed to FITC-conjugated goat anti-mouse Ig (Sigma, Paris, France) for 20 min at room temperature in the dark, and then washed and resuspended in 500 µl HBSS. Analysis was performed on a FACScan (Becton Dickinson), with the channel number (log scale) representing the mean fluorescence intensity of 10,000 cells.

PMN apoptosis

DNA fragmentation: DNA was isolated from 10⁷ control PMN and PMN from patients who had asymptomatic HIV infection (group 1) at 8 and 16 hours of transmigration as well as from control and HIV PMN in HBSS at 37° C at the same time points. PMN DNA fragmentation was conducted according to the procedure for assaying DNA fragmentation in total genomic DNA. In brief, the cells were lysed with TES buffer (20 mM Tris HCL, 200 mM EDTA, and 1% SDS) with RNAse (20 µg/ml; Boehringer Mannheim, Indianapolis, IN) at 37° C for 1 hour. Proteins were denaturated by incubation with proteinase K (1 mg/ml; Boerhinger Mannheim) at 55° C for 3 hours. The denatured protein was removed by phenol extraction. The DNA was then precipitated with alcohol overnight at ­ 20° C. The next day the DNA was rinsed with alcohol, mixed with loading buffer, and then electrophoresed in a 2% agarose gel containing 10 µg/ml ethidium bromide. The gel was examined and photographed under UV light to detect regular DNA fragmentation pattern (laddering) characteristic of apoptosis.

Morphological study: the morphological changes of apoptosis were also investigated in both transmigrated control PMN and HIV PMN by light microscopic examination of Wright-stained cytospins.

Statistical analysis

Myeloperoxidase and flow cytometric assays were compared by Student's t test. Values are expressed as the mean and SEM of "n" number of experiments.

RESULTS

Impairment of HIV PMN transmigration induced by f-MLP or IL-8 across T84 cell monolayers

The mean PMN transepithelial migration was lower for patients who suffered from asymptomatic HIV infection (Group 1) compared to healthy donors (15.1 + 1.5 versus 23.2 ± 2.6 x 10⁴ PMN CE transmigrating at 10^­7M f-MLP, respectively for patient PMN versus control PMN, n: 45, p < 0.01, Figure 1A). Decreased PMN transepithelial migration was also detectable in the asymptomatic HIV population when lower concentrations (10^­8 and 10^­9 M) of f-MLP was used to trigger neutrophil chemotaxis (Figure 1A) or when PMN transepithelial migration was induced by IL-8 (10.2 ± 2 versus 15.1 ± 1.5 x 10⁴ PMN CE transmigrating at 100 ng/ml IL-8, respectively for patient PMN versus control PMN, n: 45, p < 0.01, Figure 1A). A more pronounced decrease was observed in AIDS-defining disease as shown in figure 1B. Thus, the mean PMN transepithelial migration was lower for patients who had AIDS (Group 2) than for control subjects (9.1 ± 1.9 versus 25.2 ± 3.1 x 10⁴ PMN CE transmigrating at 10^­7 M f-MLP, respectively for patient PMN versus control PMN, n: 18, p < 0.001, Figure 1B). Significant differences were also observed at 10^­8 M f-MLP (Figure 1B) or when PMN transepithelial migration was induced by IL-8 (4.2 ± 1.2 versus 12.4 ± 1.5 x 10⁴ PMN CE transmigrating at 100 ng/ml IL-8, respectively for patient PMN vs control PMN, n: 18, p < 0.001, Figure 1B). These differences were noted irrespective of risk factor, treatment, age and sex of the patients (data not shown).

Immunoelectron study of CD11b antigen expression in control and HIV PMN

Immunoelectron microscopy analysis of anti-CD11b antibody-labeled PMN revealed numerous beads located at the plasma membrane of the resting cells. The number of beads was not significantly different in groups 1 and 2 and in control PMN (79 ± 7 versus 72 ± 10 versus 88 ± 9, repectively for group 1 versus group 2 versus control) (Figure 2A-C). A few beads were also noted inside the cytoplasm. Stimulation by f-MLP greatly increased the number of beads bound at the cell surface. No significant differences in the intensity of labeling were visible among the PMN collected from the different groups (167 ± 17 versus 159 ± 10 versus 172 ± 12, repectively for group 1 versus group 2 versus control, the differences are not significant) (Figure 2D-F).

Flow cytometry expression of CD11b and CD47 molecules in control and HIV PMN

C11b and CD47 expression was first studied on resting PMN and after stimulation with f-MLP (10^­7 M, 2 hours) and compared with the expression of these molecules after 2 hours of transmigration. As shown in Figure 3, the median fluorescence intensity of FITC-CD11b antibody bound on resting PMN was identical in groups 1 and 2 and in the healthy donors (Figure 3A-C). After stimulation with f-MLP, in the absence of migration, median fluorescence intensity increased but remained identical among groups 1 and 2 and in the control group (Figure 3A-C). Expression of CD11b after transmigration was markedly increased in group 1 and control PMN, but at similar levels (Figure 4A and B). However, this parameter could not be assessed in group 2 because of the limited number of PMN crossing the T84 monolayers. When resting and f-MLP-stimulated PMN were labeled with CD47 antibody, the median fluorescence intensity was identical in groups 1 and 2 and was slightly increased after f-MLP stimulation in the control group (Figure 3D-F). Even though, expression of CD11b and of CD47 after transmigration was increased in group 1 and control PMN, a non-significant difference was observed in these 2 populations (Figure 4A and B and 4C and D) respectively. Expression of CD47 antigen could not be evaluated on transmigrated PMN collected from group 2 donors.

Apoptosis of post-transmigrated control and HIV PMN

Electrophoresis of DNA isolated from post-transmigrated control and HIV PMN showed that DNA fragmentation (laddering bands at 200-bp intervals), a hallmark of apoptosis, was visible after 16 hours of migration, while it was undetectable after 8 hours of migration. However, no difference could be shown between control and HIV PMN (Figure 5). This was confirmed by morphological changes, i.e. decreased cytoplasm and pyknotic nuclei, as assessed by light microscopy after 16 hours of migration (data not shown).

DISCUSSION

Recurrent intestinal infections associated with diarrhea are commonly observed in AIDS patients, as well as in HIV patients presenting normal levels of CD4⁺ lymphocytes. The cause of this defect remains to be elucidated. In an attempt to address this question we investigated a possible defect in PMN chemotaxis by assessing this migration through a monolayer of intestinal epithelial cells in an in vitro system. We found that migration of HIV PMN across intestinal epithelial cells was significantly depressed. This decreased migration was more pronounced in HIV PMN isolated from patients with CD4⁺ lymphocyte counts < 200/mm³, (group 2), but was also observed in patients who had asymptomatic disease and CD4⁺ lymphocyte counts > 200/mm³, (group 1). This study provides the first evidence of the impairment of HIV neutrophil migration through an intestinal epithelial barrier. To date, disparate data have been accumulated on the functionality of PMN isolated from HIV-infected patients. Defects of phagocytosis, antimicrobial activity and superoxide production have been reported [2-4, 6]. Normal or increased phagocytosis and normal or elevated production of toxic oxygen radicals despite diminished bactericidal activity have also been reported [1, 5, 8]. Some investigators have observed a normal chemotaxis in PMN from HIV patients or impaired chemotaxis in early asymptomatic disease followed by normalization with disease progression [4, 30, 31]. In other studies of HIV-infected patients, a chemotaxis defect of PMN has been observed [32, 33]. However, these studies were performed on isolated cells or in suspension. To the best of our knowledge, this present work is the first study aimed at assessing chemotaxis function in HIV PMN using an in vitro model requiring cell-cell interactions between PMN and epithelial cells.

The discrepancies between studies on HIV PMN functions may be attributed in part to the diversity of experimental procedures used to measure PMN functions. Utilisation of autologous serum may influence the results, since it has been shown that elevated levels of cytokines, such as TNF-alpha decrease PMN functions [34-36]. In some studies of HIV-infected patients, the most pronounced reduction of PMN functions has been observed in patients with low CD4⁺ counts. Acute viral and bacterial infections may also significantly influence PMN function [37]. Antimicrobial agents and narcotic drugs may influence PMN functions and such agents are frequently used by HIV-infected patients [38, 39]. In the present work, transepithelial migration of PMN was carried out in the absence of autologous serum, in order to determine if impairment in neutrophil migration stemmed from intrinsic defects. In addition, it is noteworthy that the HIV PMN samples collected in our study exhibited a decreased transepithelial migration, regardless of variable risk factors, drug treatments and/or concurrent infections.

Different factors may influence neutrophil transepithelial migration. Patients with leukocyte adhesion deficiency (LAD) syndrome present a primary defect in the common ß subunit of Mac-1, LFA-1, and p150,95 proteins that produce recurrent bacterial intestinal infections [40]. This is confirmed by in vitro studies showing that PMN from patients with LAD fail to elicit a transepithelial migration [26]. The defect however may be intracellular as substantiated by a recent report of a patient with clinical features of congenital LAD-1 expressing normal levels of CD11b/CD18 in PMN. In this patient, stimuli failed to induce a conformational change of CD11b/CD18 integrin receptors that normally switches their ligand binding capacity from a low to a high-avidity state. Consequently, in vitro chemotaxis and endothelial transmigration of these neutrophils were found to be almost absent [41]. Neutrophil passage across the T84 cells is reported to require two steps. First, PMN have to adhere to the epithelial cells, which is mediated by CD11b/CD18 [26]. Afterwards, the PMN have to progress between epithelial cells via a paracellular pathway that involves CD47 [27]. We thus analyzed, by flow cytometry, the expression of CD11b, CD18 and CD47 at the surface of HIV PMN before and after transepithelial migration. We failed to detect any significant difference in the expression of these antigens between control and HIV PMN. In particular, the functional up-regulation seen upon PMN activation during the transmigration and/or after f-MLP stimulation was the same in control PMN and in HIV-infected PMN. However, as discussed above, a nonfunctional assembly of the CD11b and CD18 subunits at the cell surface might lead to a diminished transepithelial migration. Upregulation of CD11b/CD18 is accompanied by the recruitment of additional alpha and beta subunits from the cytoplasm [3]. Given that PMN can be infected by HIV [42, 43], one can assume that the virus could interfere with the cytoskeletal architecture, leading to a nonfunctional assembly of CD11b and CD18 subunits [44-46]. Another possible explanation is defective chemoattractant receptors. However this is unlikely since PMN transepithelial migration induced by both f-MLP and IL-8 is decreased in our study. The recent identification of certain chemokine receptors (CCR5 and CXCR4) as coreceptors for HIV, has provided tremendous insight into the mechanisms underlying viral entry and tropism, and it has been shown that HIV proteins such as gp120 could mask these cell surface receptors [47]. Although f-MLP and IL8 receptors have not been yet implicated in HIV cell entry and are structurally very distinct from CXR4 receptors, the hypothesis that some HIV proteins could hinder f-MLP and IL-8 surface receptors cannot be eliminated. An alternative explanation would be that the level of receptors for fMLP or IL-8 are down-regulated at the cell surface in HIV patients. More work is needed to clarify this point. A defective transduction of the signal mediated by the chemoattractant could be also envisaged to explain the impairment of HIV PMN migration. In our study the defect in PMN transepithelial migration was more evident in patients with low counts of CD4⁺ lymphocytes, suggesting that abnormalities in the signal transduction and/or cytoskeletal organization are accentuated in later stages of the disease. Another motility disorder, named neutrophil actin dysfunction has been described [48], which is an inherited disease where the PMN show profound motility abnormalities, abnormal actin polymerization and impairment of actin assembly. In AIDS patients, further studies are needed to clarify whether HIV infection might interfere with cytoskeletal organization of microfilaments.

Because apoptotic PMN are functionally impaired in AIDS patients, it has been hypothesized that the abnormalities of PMN function observed in the course of HIV infection might be the result of an accelerated apoptosis [7]. However we failed to detect any difference in the spontaneous apoptotic process between transmigrated control and HIV PMN.

CONCLUSION

REFERENCES

1. Bandres J C, Trial J, Musher D M, Rossen R D. 1993. Increased phagocytosis and generation of reactive oxygen products by neutrophils and monocytes of men with stage I human immunodeficiency virus infection. J. Infect. Dis. 168: 75.

2. Chen T P, Roberts R L, Wu K G, Ank B J, Stiehm E R. 1993. Decreased superoxide anion and hydrogen peroxide production by neutrophils and monocytes in human immunodeficiency virus-infected children and adults. Pediatr. Res. 34: 544.

3. Elbim C, Prevot MH, Bouscarat F, Franzini E, Chollet-Martin S, Hakim J, Gougerot-Pocidalo M A. 1994. Polymorphonuclear neutrophils from human immunodeficiency virus-infected patients show enhanced activation, diminished fMLP-induced L-selectin shedding, and an impaired oxidative burst after cytokine priming. Blood 84: 2759.

4. Flo R W, Naess A, Nilsen A, Harthug S, Solberg C O. 1994. A longitudinal study of phagocyte function in HIV-infected patients. AIDS 8: 771.

5. Jarstrand C, Akerlund B. 1994. Oxygen radical release by neutrophils of HIV-infected patients. (Review) Chem. Biol. Interact. 91: 141.

6. Pitrak D L, Bak P M, De Marais P, Novak R M, Andersen B R. 1993. Depressed neutrophil superoxide production in human immunodeficiency virus infection. J. Infect. Dis. 167: 1406.

7. Pitrak D L, Tsai H C, Mullane K M, Sutton S H, Stevens P. 1996. Accelerated neutrophil apoptosis in the acquired immunodeficiency syndrome. J. Clin. Invest. 98: 2714.

8. Ryder M I, Winkler J R, Weinreb R N. 1988. Elevated phagocytosis, oxidative burst and F-actin formation in PMN from individuals with intraoral manifestation of HIV infection. J. Acquir. Immune. Defic. Syndr. 1: 346.

9. Spear G T, Ou C Y, Kessler H A, Moore J L, Schochetman G, Landay A L. 1990. Analysis of lymphocytes, monocytes, and neutrophils from human immunodeficiency virus (HIV)-infected persons for HIV DNA. J. Infect. Dis. 162: 1239.

10. Martin L S, Spira T J, Orloff S L, Holman R C. 1988. Comparison of methods for assessing chemotaxis of monocytes and polymorphonuclear leukocytes isolated from patients with AIDS or AIDS-related conditions. J. Leukocyte Biol. 44: 361.

11. Sewankambo N, Mugerwa R D, Goodmame R. Carswell J W, Moody A, Lloyd G, Lucas S B. 1987. Enteropathic AIDS in Uganda. An endoscopic, histological, and microbiological study. AIDS 1: 9.

12. Smith P D, Lane H C, Gill V J. Manisschewitz J F, Quinnan G V, Fauci A S, Masur H. 1988. Intestinal infections in patients with the acquired immunodeficiency syndrome (AIDS). Etiology and response to therapy. Ann. Intern. Med. 108: 328.

13. Gillin J S, Shike M, Alcock N, Urmacher C, Krown S, Kurtz R C, Lightdale C J, Winawer S J. 1985. Malabsorption and mucosal abnormalities of the small intestine in the acquired immunodeficiency syndrome. Ann. Intern. Med. 102: 619.

14. Kotler D P, Gaetz H P, Klein E B, Lange M, Holt P R. 1984. Enteropathy associated with the acquired immunodeficiency syndrome. Ann. Intern. Med. 101: 421.

15. Celum C L, Chaisson R E, Rutherford G W, Barnhart L, Echenberg D F. 1987. Incidence of salmonellosis in patients with AIDS. J. Infect. Dis. 156: 998.

16. Kotler D P, Glang T T, Thiim M, Nataro J P, Sordillo E M, Orenstein J M. 1995. Chronic bacterial enteropathy in patients with AIDS. J. Infect. Dis. 171: 552.

17. Nelson M R, Shanson D C, Hawkins D A, Gazzard B G. 1992. Salmonella, Campylobacter and Shigella in HIV-seropositive patients. AIDS 6: 1495.

18. Nichols L, Balogh K, Silverman M. 1989. Bacterial infections in the acquired immune deficiency syndrome. Clinicopathologic correlations in a series of autopsy cases. Am. J. Clin. Pathol. 92: 787.

19. Kotler D P, Reka S, Borcich A, Cronin W J. 1991. Detection, localization and quantitation of HIV-associated antigens in intestinal biopsies from HIV-infected patients. Am. J. Pathol. 139: 823.

20. Ullrich R, Zeitz M, Heise W, L'age M, Hoffken G, Rieken E O. 1989. Small intestinal structure and function in patients infected with human immunodeficiency virus (HIV): evidence for HIV-induced enteropathy. Ann. Intern. Med. 111: 15.

21. Kotler D P, Reka S, Clayton F. 1993. Intestinal mucosal inflammation associated with human immunodeficiency virus infection. Dig. Dis. Sci. 38: 1119.

22. Yahi N, Baghdiguian S, Bolmont C, Fantini J. 1992. Replication and apical budding of HIV-I in mucous-secreting colonic epithelial cells. J. Acquir. Immune Defic. Syndr. 5: 993.

23. Madara J L, Colgan S, Nusrat A, Delp C, Parkos C. 1992. A simple approach to measurement of electrical parameters of cultured epithelial monolayers: use in assessing neutrophil-epithelial monolayers. J. Tiss. Cult. Meth. 14: 209.

24. Madara J L. Pathobiology of the intestinal epithelial barrier. 1990. (Review) Am. J. Pathol. 137: 1273.

25. Arnaout M A. 1990. Structure and function of the leukocytes adhesion molecules CD11/CD18. (Review) Blood 75: 1037.

26. Parkos C A, Delp C, Arnaout M A, Madara J L. 1991. Neutrophil migration across a cultured intestinal epithelium. Dependence on a CD11b/CD18-mediated event and enhanced efficiency in physiological direction. J. Clin. Invest. 88: 1605.

27. Parkos C A, Colgan S P, Liang T W, Nusrat A, Bacarra A E, Carnes D K, Madara J L. 1996. CD47 mediates post-adhesive events required for neutrophil migration across polarized intestinal epithelia. J. Cell Biol. 132: 437.

28. Lindberg F P, Bullard D C, Caver T E, Gresham H D, Beaudet A L, Brown E J. 1996. Decreased resistance to bacterial infection and granulocyte defects in IAP-deficient mice. Science 274: 795.

29. Anonymous. 1992. Centers for diseases control. 1993 revised classification system for HIV infection and expanded surveillance case definition for AIDS among adolescents and adults. Morb. Mortal Wkly. Rep. 41: 1.

30. Nielsen H, Kharazmi A, Faber V. 1986. Blood monocyte and neutrophil functions in the acquired immune deficiency syndrome. Scand. J. Immunol. 24: 291.

31. Roilides E, Mertins S, Eddy J, Walsh T J, Pizzo P A, Rubin M. 1990. Impairment of neutrophil chemotactic and bactericidal function in children infected with human immunodeficiency virus type 1 and partial reversal after in vitro exposure to granulocyte-macrophage colony stimulating factor. J. Pediatr. 117: 531.

32. Ellis M, Gupta S, Galant S, Hakim S, Vandvev C, Toy C, Cairo M S. 1988. Impaired neutrophil function in patients with AIDS or AIDS related complex: a comprehensive evaluation. J. Infect. Dis. 158: 1268.

33. Lazzarin A, Uberti-Foppa C, Galli M, Mantovani A, Poli G, Franzetti F, Novati R. 1986. Impairment of polymorphonuclear leucocyte function in patients with acquired immunodeficiency syndrome and with lymphadenopathy syndrome. Clin. Exp. Immunol. 65: 105.

34. Kapp A, Komann A, Schopf E. 1991. Effect of tumor necrosis factor alpha in vivo on human granulocyte oxidative metabolism. Arch. Dermatol. Res. 283: 362.

35. Pinegin B V, Saidov M Z, Scheltsyna T L, Gabrilovich D I, Khaitov R M. 1993. Peripheral blood neutrophils from HIV-infected individuals are armed with factors that cause inhibition of their migration in response to specific antigens. Immunol. Lett. 36: 13.

36. Gabrilovich D I, Aveeva L A, Serebrovskaya L V, Shepeleva G K, Suvorova Z K, Buravcova E V, Oganezov V K, Platonov A E, Habarova V V, Posly I M, Pokrovsky V V. 1993. Impact of HIV-positive sera on the functional activity of polymorphonuclear neutrophils from healthy donors. Scand. J. Immunol. 37: 159.

37. Solberg C O, Kalager T, Hill H R, Glette J. 1982. Polymorphonuclear leukocyte function in bacterial and viral infections. Scand. J. Infect. Dis. 14: 11.

38. Oleske J M, de la Cruz A, Ahdieh H. 1983. Effects of antibiotics on polymorphonuclear leukocyte cheminescence and chemotaxis. J. Antimicrob. Chemother. 12: S35.

39. Tubaro E, Avico U, Santiageli C, Zuccaro P, Cavallo G, Pacifici R, Croce C, Borelli G. 1985. Morphine and methadone impact on human phagocytic physiology. Int. J. Immunopharmacol. 7: 865.

40. Gresham H D, Graham I L, Anderson D C, Brown E J. 1991. Leukocyte adhesion-deficient neutrophils fail to amplify phagocytic function in response to stimulation. Evidence for CD11b/CD18 dependent and independent mechanisms of phagocytosis. J. Clin. Invest. 88: 588.

41. Kuijpers T W, Van Lier R A, Hamann D, de Boer M, Thung L Y, Weening R S, Verhoeven A J, Roos D. 1997. Leukocyte adhesion deficiency type 1 (LAD-1)/variant. A novel immunodeficiency syndrome characterized by dysfunctional ß2 integrins. J. Clin. Invest. 100: 1725.

42. Gabrilovich D I, Vassilev V, Nosikov V V, Serebrovskaya L V, Ivanova L A, Pokrovsky V V. 1993. Clinical significance of HIV DNA in polymorphonuclear neutrophils from patients with HIV infection. J. Acquir. Immune Defic. Syndr. 6: 587.

43. Kojouharoff G, Ottman O G, von Briesen H, Geissler G, Rubsamen-Waigmann H, Hoelzer D, Ganser A. 1991. Infection of granulocyte/monocyte progenitor cells with HIV-1. Res. Virol. 142: 151.

44. Choudhury S, el-Farrash M A, Kuroda M J, Harada S. 1996. Retention of HIV-1 inside infected MOLT-4 cells in association with adhesion-induced cytoskeleton reorganization. AIDS 10: 363.

45. Luftig R B, Lupo L D. 1994. Viral interactions with the host-cell cytoskeleton: the role of retroviral proteases. Trends Microbiol. 2: 178.

46. Rey O, Canon J, Krogstad P. 1996. HIV-1 Gag protein associates with F-actin present in microfilaments. Virology 220: 530.

47. Tarasova N I, Stauber R H, Michejda C J. 1998. Spontaneous and ligand-induced trafficking of CXC-chemokine receptor 4. J. Biol. Chem. 273: 15883.

48. Boxer L A, Hedley-Whyte E T, Stossel T P. 1974. Neutrophil actin dysfunction and abnormal neutrophil behavior. N. Engl. J. Med. 291: 1093.