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(Journal of Leukocyte Biology. 2002;72:650-656.)
© 2002 by Society for Leukocyte Biology

Defective migration of monocyte-derived dendritic cells in LAD-1 immunodeficiency

Maurilia Fiorini*,{dagger}, William Vermi{ddagger}, Fabio Facchetti{ddagger}, Daniele Moratto*,{dagger}, Giulio Alessandri§, Lucia Notarangelo*,{dagger}, Arnaldo Caruso§, Piergiovanni Grigolato{ddagger}, Alberto G. Ugazio*,{dagger}, Luigi D. Notarangelo*,{dagger} and Raffaele Badolato*,{dagger}

* Istituto di Medicina Molecolare "Angelo Nocivelli",
{dagger} Clinica Pediatrica,
{ddagger} Cattedra di Anatomia Patologica, and
§ Istituto di Microbiologia, Università di Brescia, Italy

Correspondence: Raffaele Badolato, M.D., Ph.D., Clinica Pediatrica, Università di Brescia, c/o Spedali Civili, 25123 Brescia, Italy. E-mail: badolato{at}master.cci.unibs.it


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
ß2 Integrins (CD18) are required for leukocyte migration. In fact, the absence of CD18 results in type-1 leukocyte adhesion deficiency (LAD-1). We analyzed the distribution phenotype and function of dendritic cells (DCs) in three LAD-1 patients with homozygous mutations of CD18. Two of them did not express CD18 (Patients A and C), and the other subject (Patient B) displayed reduced expression of ß2 integrins because of a missense mutation. Analysis of DCs derived from Patients A and B showed an abnormal morphology and a severe impairment in transendothelial migration and chemotactic response to CCL19/macrophage inflammatory protein-3ß, suggesting that CD18 is required for migration of monocyte-derived DCs. Nevertheless, DCs displayed normal macropinocytosis and underwent normal maturation after addition of tumor necrosis factor {alpha}. Finally, immunohistochemical analysis of lymph nodes from subjects B and C revealed a significant reduction in the number of factor-XIIIa+ interstitial DCs in the interfollicular area in both patients, suggesting that CD18 plays a role in the migration of these cells in vivo.

Key Words: human • immunodeficiency diseases • adhesion molecules • cell trafficking


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Type-1 leukocyte adhesion deficiency (LAD-1) is a rare, autosomal recessive disorder characterized by delayed separation of umbilical cord, recurrent bacterial infections of the skin, and mucosal membranes without pus formation, despite persistent neutrophilia (up to 100,000 cells/mm3) [1 , 2 ]. The underlying molecular defect has been shown to reside in the gene encoding the ß2 integrin chain, a 95-kD glycoprotein designed as leukocyte antigen CD18, which is non-covalently associated with three distinct {alpha} chains that have a leukocyte antigen designation of CD11a/CD18 (lymphocyte function-associated antigen-1), CD11b/CD18 (Mac-1), and CD11c/CD18 (p150,95) [3 ]. These glycoproteins are expressed on the cell surface of leukocytes and mediate important leukocyte functions, including adhesion, transendothelial migration, and phagocytosis. The recognized immunological defect of LAD-1 consists of a severe impairment in neutrophil emigration to tissue, as the CD11/CD18 heterodimers are pivotal for firm adhesion and transendothelial migration of these circulating cells [3 ]. CD18 is expressed at high levels in dendritic cells (DCs) and is required for migration of these cells [4 , 5 ]. Studies of DCs in CD18 null mice have demonstrated a significant reduction in the number of interstitial DCs in the lungs of these animals, suggesting that expression of CD18 is required for migration of this DC subset [6 ].

DCs represent a heterogeneous population, including blood precursors, immature DCs in the peripheral tissue, and terminally matured cells in secondary lymphoid organs. Immature DCs are found throughout the body, but are present in large number in areas of potential antigen entry where they exert a sentinel function [7 , 8 ]. They are highly specialized in antigen capture and processing by means of surface receptors such as the Fc portion of immunoglobulins (Ig) and mannose receptor [9 ]. On the basis of their phenotype and functional property, CD11c+ myeloid DCs are distinct in two population, Langerhans cells (LC) and interstitial DCs. LC are a subset of DCs that express CD1 and Langerin and are located at the epithelial surface of the skin, gastrointestinal, respiratory, and genitourinary tracts [10 11 12 ]. In contrast, interstitial DCs are recognizable by intracellular expression of factor XIIIa and are located in the dermis (dermal DCs), in the interstitial areas of solid organs, and, admixed with lymphocytes, in the interfollicular area of the lymph nodes [13 14 15 ]. Like other leukocytes, DCs need to tether to the endothelium, achieve a firm adhesion, and finally extravasate and migrate within the tissue and secondary lymphoid organs through a process that involves interaction of CD18 with intercellular adhesion molecule-1 (ICAM-1) [5 ].

Until now, studies of DC function and phenotype in patients affected with LAD-1 are completely lacking. In this paper, we report the characterization of DCs in three LAD-1 patients. Immunohistochemical analysis of lymph nodes revealed a severe depletion of factor XIIIa+ interstitial DCs in the interfollicular area. In vitro culture of peripheral blood monocytes from LAD-1 patients resulted in generation of a DC subset with normal expression of intracellular factor XIIIa+. These DCs underwent normal maturation with expression of CD83 and CD86, but displayed an abnormal morphology and a severe impairment in chemotaxis and transendothelial migration, suggesting that CD18 plays a role in migration of interstitial DCs.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Patients
Three patients with clinical and molecular features of LAD-1 were included in this study. All of them presented a severe course of disease that was recognized within 3 months of life because of repeated episodes of cutaneous infections and otitis, which resulted in sepsis. In Patients A and B, Staphylococcus aureus was isolated from cutaneous "cold abscesses," and in Patient C, otitis was sustained by Pseudomonas. All patients presented sepsis associated to bacterial skin infections and required aggressive antibiotic treatment. Leukocyte counts varied from 27,000 up to 160,000 cells/µl depending on infections. A delayed separation of the umbilical cord associated to omphalitis was recognized only in Patient A. Blood samples and lymph node specimens were obtained upon informed consent from Patients B and C and from three age-matched subjects that were hospitalized for minor head trauma.

Genetic analysis of CD18 revealed homozygous mutations in all three patients. Both parents of Patients B and C presented the same mutation of the proband in a heterozygous state. In Patient A, there was evidence of parental consanguinity. In Patient A, direct cDNA sequencing revealed a deletion of 181 bp, which corresponded to exon 4; this mutation results in a frame shift, leading to a predicted, premature termination of protein translation at codon 117 that would prevent protein expression. In Patient B, we identified a substitution of T 1906 with C that results in the change of cysteine 612 into arginine, possibly leading to conformational and/or functional changes of the ß2 integrin complex. Direct sequencing of reverse transcriptase-polymerase chain reaction (RT-PCR) fragments in Patient C revealed a 10-nucleotide deletion (from 191 to 200 bp) in exon 3 of the CD18 gene. This mutation, which results in frame shift with premature termination, had already been reported in an unrelated LAD-1 patient described by Lopez Rodriguez et al. [16 ], thereby suggesting the possibility of a common mechanism in the generation of the deletion. Patients were evaluated and treated at the Department of Pediatrics, University of Brescia, Italy.

Reagents
Human recombinant granulocyte macrophage-colony stimulating factor (GM-CSF), interleukin (IL)-4, and tumor necrosis factor {alpha} (TNF-{alpha}) were obtained from Pepro Tech (Rocky Hill, NJ). All reagents and media, tested by the endotoxin kit (Sigma Chemical Co., St. Louis, MO), contained endotoxin at concentrations below the detection levels (12 pg/ml). CD18 (clone L130), CD11a (clone G-25.2), CD11b (clone D12), and CD11c (S-HCL) were obtained from Becton Dickinson Immunocytometry Systems (San Jose, CA); control mouse IgG (clone X0943), CD14 (clone TUK4), fluorescein isothiocyanate (FITC)-conjugated goat anti-mouse IgG Fab2, and anti-factor XIIIa (clone F8/86) were from Dako (Golstrup, Denmark); and CD83 (clone HB15a) and CD86 (HA5.2B7) were purchased from Coulter-Immunotech (Marseilles, France).

Flow cytometry analysis
Analysis of cell surface expression of leukocyte antigens was performed as previously described [17 ]. Briefly, blood samples (100 µl) or purified cells (5x105 cells) were incubated with the appropriate antibodies or control mouse IgG for 30 min at 4°C. Samples were washed twice with phosphate-buffered saline (PBS), suspended in 100 µl PBS, and incubated at 4°C for 30 min with 4 µl FITC-conjugated goat anti-mouse IgG Fab2. When required, red blood cells were lysed by incubating the samples with 4 ml lysing buffer (lysing solution, Becton Dickinson). Washed cells were fixed in 1% paraformaldehide; at least 5000 cells were analyzed by FACScan (Becton Dickinson). Monocytes or DCs were gated on the basis of forward and side scatter. Intracellular staining of DCs with antifactor XIIIa was performed by the Cytofix/Cytoperm Plus kit, according to the manufacturer’s instructions (Pharmingen, San Diego, CA).

Cell culture
Peripheral blood mononuclear cells (PBMC) were purified by Ficoll separation medium (Lympholyte H, Cedarlane Laboratories, Hormby, Ontario, Canada) gradient centrifugation as described elsewhere [17 ]. When indicated, monocytes were purified by Percoll separation medium (Pharmacia Biotech, Upsala, Sweden) as described elsewhere [17 ]. Monocytes were more than 97% pure as determined by direct immunofluorescence assay using the monoclonal antibody (mAb) CD14 (Dako). Cells were cultured in RPMI 1640 (HyQ, HyClone Europe Ltd., Cramlington, UK), containing 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mmol/L glutamine, 20 mmol/L HEPES (Imperial, UK), and 10% heat-inactivated fetal calf serum (FCS; Boehringer Mannheim GmbH, Germany). DCs were generated in vitro from blood-derived monocytes as previously described [5 ]. Briefly, highly enriched monocytes were cultured for 7 days at 1 x 106 cells/ml in 24-well tissue culture plates (Nunc, Roskilde, Denmark) in RPMI 1640 with 10% FCS supplemented with GM-CSF (100 ng/ml) and IL-4 (100 ng/ml). When indicated, DCs were incubated for an additional 2 days with TNF-{alpha} (10 ng/ml). After 7 days of culture, cells generated in vitro were CD1a+ (CD1a clone O10, Coulter-Immunotech) and CD14- (Dako).

RT-PCR analysis and cDNA sequencing
Extraction of total RNA was purified using RNAzol (Tel Test Inc., Friendswood, TX) from PBMC or from Epstein-Barr virus-transformed cell lines following the manufacturer’s instructions. cDNA synthesis was performed with 1 µg RNA in a total volume of 20 µl containing 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 5 mM MgCl2, 1 mM each dNTP, 1 U/µL RNase ribonuclease inhibitor (Life Technologies, Gaithersburg, MD), 2.5 µM random hexamers, and 2.5 U/µL MuLV RT (Perkin Elmer, Foster City, CA). The amplified cDNA fragments were sequenced by using the dRhodamine Terminator Cycle sequencing kit (PE Applied Biosystems, Warrington, UK) and automated Abi Prism 310 DNA sequencer.

Immunohistochemistry
Inguinal lymph node biopsies were obtained from Patients B and C at the age of 4 and 5 months, respectively. All specimens were fixed in formalin and embedded in paraffin; in addition, half specimen was also frozen in liquid nitrogen and stored at 80°C. Immunohistochemistry was performed with an indirect streptavidin biotin complex immunoperoxidase technique on paraffin or cryostat sections. Quantitative analysis of interstitial DC numbers was performed by counting three high power fields (HPF) in a blind evaluation after coding samples. Interstitial factor XIIIa+ DCs were identified on the basis of cytoplasmic staining and appropriate morphology. The following antibodies were applied: CD3 (clone SK7), CD11a, CD11b, and CD11c (Becton Dickinson); CD18 (clone MEM48, Ylem, Rome, Italy); CD20 (clone B-Ly1) and CD45R0 (clone UCHL1, Dako); CD83, DC-LAMP, and Langerin (clone DCGM4, Coulter-Immunotech); and factor XIIIa (Dako).

Migration assays
Migration of DCs [5x10; 5 cells/ml in RPMI 1640+1% bovine serum albumin (BSA)] was evaluated by a microchamber technique as described elsewhere [17 ] by using 5-µm pore size polycarbonate filters (Neuro Probe, Gaithersburg, MD). At the end of the incubation (120 min), filters were removed, fixed, and stained by Diff-Quick (Harleco, Gibbstown, NJ), and six oil immersion fields were counted in a blind evaluation after coding samples.

Transendothelial migration
Primary cultures of human adrenal gland capillary endothelial cells (EC) were obtained as previously described [18 ]. EC (5x104) were seeded on collagen-coated trans-well culture inserts (6.5-mm diameter polycarbonate clear membrane with 3-µm pores; Costar Corp., Cambridge, MA). EC were cultured in EBM complete medium until confluence was reached. Under these conditions, EC did not cross the membrane and formed a complete monolayer, usually within 2 days of culture and only on the upper surface of the filter, as confirmed by staining a batch of trans-well inserts with Diff-Quick before their use in the experiments. The EC monolayer was treated for 4 h with 5 ng/ml TNF-{alpha} or with EBM complete medium alone. The cells were washed three times with RPMI 1640 containing 0.25% BSA. RPMI-1640 complete medium (0.6 ml) was added to each well of a 24-well plate. DCs (2x105 cells in 200 µl complete RPMI-1640 medium) were added into the insert before the immersion of the trans-well. The 24-well plates were incubated for 18 h at 37°C in a 5% CO2 atmosphere; previous experiments of TEM of DCs at different time points such as 6, 12, 18, and 24 h showed that the highest number of migrated cells was reached at 18 h. After this incubation time, the trans-well inserts were then removed, and cells transmigrated into the well were collected by centrifugation and suspended in PBS. By an optic microscope, the cells contained in three HPF (40x) were counted. The tests were run in triplicates.

Statistical analysis
Comparison between normal donors and LAD-1 patients was performed where indicated by nonparametrical tests.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Expression of CD18 protein in LAD-1 patients
Diagnosis of LAD-1 was established on the basis of a profound defect of CD18 expression as determined by flow cytometry. As shown in Figure 1A , CD18 was not detectable on the monocyte cell surface of Subjects A and C, and low levels of CD18 expression were detectable in Patient B. Other leukocyte subsets did not display detectable levels of CD18 or the ß2-associated proteins CD11a, CD11b, and CD11c (data not shown). The subject with a partial CD18 expression, Patient B, presented a milder course of the disease, although he had two episodes of sepsis at 1 and 3 months of life.



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  		Figure 1. CD18 expression in peripheral blood and lymph nodes of LAD-1 patients. (A) Flow cytometry analysis of CD18 expression. PBMC derived from LAD-1 patients (Patients A, B, and C) were incubated with control mouse-IgG (thin line) or FITC-conjugated CD18 (thick line) mAb. After washing, cells were fixed with paraformaldehide and analyzed by a flow cytometer. Monocytes were gated on the basis of forward and side scatter, and green fluorescence of at least 5000 events was analyzed. The x-axis represents the intensity of green fluorescence expressed in a log scale as mean channel, and the y-axis represents the relative cell number. (B) Expression of CD18 in lymph nodes. In a control lymph node, the majority of leucocytes strongly expresses CD18 protein. Lymph node section from Patient B contains a population of CD18+ cells in the paracortex. In contrast, no positivity is detectable in Patient C.

 
Analysis of ß2 integrin expression by immunohistochemistry in the lymph node from Patient B revealed numerous CD18+ cells, and the positivity was more evident on macrophages and interdigitating DCs (IDC; Fig. 1B ). In contrast, analysis of the lymph node of Patient C demonstrated complete absence of CD18, as compared with a reactive lymph node obtained from a normal subject (Fig. 1B) . In addition, CD11a and CD11c were noticed on scattered cells including IDC, and CD11b was totally absent (data not shown).

Lymph nodes from LAD-1 patients are depleted of factor XIIIa+ interstitial DCs
Lack of CD18 from the cell surface may affect the recruitment of various leukocyte subsets to the lymph node, resulting in depletion of lymphoid tissue [19 ]. On histology, lymph nodes from Patients B and C were small (maximum diameter, 5 mm). The parenchyma was depleted of lymphoid cells: the cortex contained small, primary CD20+ B follicles (Fig. 2a 2b 2c ), whereas the architectural structure of paracortex was severely altered (Fig. 2d 2e 2f) .



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  		Figure 2. Depletion of lymphocytes and factor XIIIa+ DCs in lymph nodes from LAD-1 patients. Serial sections of lymph nodes from a control (a, d, g, l), Patient B (b, e, h, m), and Patient C (c, f, i, n) stained, respectively, for anti-CD20 (a–c) and anti-CD3 (d–f). In the control, a full-blown lymphoid follicle (a) is surrounded by a normal T cell population (d). In Patients C and B, a depletion of CD20+ (b, c) and CD3+ (e, f) lymphocytes is evident. A reduction of factor XIIIa+ DCs can be observed in Patients B (h) and C (e) in comparison with the control (g). In contrast, the paracortical T cell area contains numerous DC-LAMP+-interdigitating DCs with similar distribution in the control (l) and in both patients (m, n).

 
Staining of lymph nodes from Patients B and C with antifactor XIIIa mAb, which identifies DCs immigrated to the lymph node from peripheral tissues or resident in the interfollicular area of lymph nodes [14 , 20 ], showed that the number of factor XIIIa+ cells was significantly reduced as compared with reactive lymph nodes (46±8 vs. 11±4 for Patient B; 7±5 for Patient C; P<0.05; Fig. 2g 2h 2i ). In contrast, the paracortex showed accumulation of numerous CD1a+ LC/IDC admixed with sparse CD3+/CD45R0+ T cells. The IDC showed strong expression of the maturation-associated antigens DC-LAMP (Fig. 2l 2m 2n) and CD83 (data not shown). Sparse Langerin cells, corresponding to recently immigrated LC, were also recognizable (data not shown).

Abnormal morphology but normal differentiation of monocyte-derived DCs in LAD-1 patients
Upon appropriate stimulation, monocytes that exit the bloodstream differentiate into professional antigen-presenting cells (APC) such as macrophages or DCs. In vitro, monocytes can be induced to differentiate into functional DCs when cultured in the presence of IL-4 and GM-CSF. We analyzed whether monocytes obtained from LAD-1 subjects A and B could differentiate into DCs upon culture with IL-4 and GM-CSF followed by addition of TNF-{alpha} in the last 2 days. To rule out the possibility that DCs derived from LAD-1 patients were impaired in the expression of the intracellular antigen factor XIIIa, we investigated the expression of this antigen in monocyte-derived DCs by intracellular staining. Flow cytometric analysis of factor XIIIa expression demonstrates that monocyte-derived DCs generated from LAD-1 patients displayed normal levels of the antigen (Fig. 3 and data not shown). Microphotographs of monocyte-derived DCs show a homogeneous expression of factor XIIIa antigen in the cytosol, suggesting that DCs from LAD-1 acquire the antigen during differentiation in vitro (Fig. 3 , insert). Parallel analysis of cell surface expression of DC-SIGN, a C-lectin that is used for rolling and migration through endothelial monolayers and interacts at high affinity with ICAM-2 and ICAM-3 antigens [21 , 22 ]—CD86, and CD83 of monocyte-derived DCs obtained from LAD-1 patients revealed normal maturation of these cells (Fig. 3) . Next, we incubated DCs generated in vitro with FITC-conjugated dextran particles for 1 h at 37°C and analyzed cellular uptake by measuring the increase of green fluorescence. As shown in Figure 3 , DCs derived from Patients A and B displayed a normal macropinocytosis as compared with a normal subject.



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  		Figure 3. Differentiation of DCs in LAD-1 patients. Monocyte-derived DCs derived from LAD-1 Patients A and B or from normal subject sections were stained with control mouse-IgG, DC-SIGN, CD86, or factor XIIIa mAb or were analyzed for macropinocytosis of FITC-dextran. The insert displays a microphotograph of intracellular staining of factor XIIIa+ cells. After incubation with goat anti-mouse-FITC, cells were washed twice, fixed with PBS 1% paraformaldehide, and analyzed by a flow cytometer. The extent of expression is presented on the y-axis as the mean of green fluorescence mean channel. Data shown are representative of two independent experiments performed on Patients A and B and a normal subject.

 
Addition of TNF-{alpha} to DC culture results in typical stellate morphology characterized by pronounced protrusions and microvillous projections of their plasmamembrane [23 ]. Figure 4 demonstrates that cells derived from Patients A and B displayed an abnormal morphology, as cells lacked cellular projections, which were present in cells derived from two normal subjects, suggesting that CD18 is important for the morphology of DCs.



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  		Figure 4. Cellular morphology of DCs from LAD-1 patients. Monocyte-derived DCs were incubated for an additional 48 h with TNF-{alpha} (10 ng/ml) and photographed using an inverted optic microscope; original scale bar, 10 µm. The results shown are microphotographs representative of two independent experiments performed on Patients A and B and two normal subject.

 
Impaired transendothelial migration of monocyte-derived DCs in LAD-1 patients
Our observation that factor XIIIa+ DCs are present in reduced number in lymph nodes from LAD-1 patients suggests that cellular motility of DCs derived from LAD-1 patients might be impaired. To investigate this hypothesis, we assessed the chemotactic response of DCs of LAD-1 Patients A and B to CCL19/macrophage inflammatory protein-3B (MIP-3B) by using a modified Boyden microchamber. As shown in Figure 5A , cells derived from both patients displayed a severe impairment in chemotactic response to stimuli as compared with normal cells. Moreover, we analyzed transendothelial, spontaneous migration of monocyte-derived DCs from two LAD-1 patients (Pt A and Pt B) and two normal subjects. We observed that an extremely low number of DCs (up to 15 cells/three HPF in Patient A; up to 40 cells/three HPF in Patient B) derived from LAD-1 patients migrated through TNF-{alpha}-activated endothelial monolayers as compared with normal subjects, suggesting that CD18 is critical for migration of monocyte-derived DCs (Fig. 5A) . Analysis of migration at other time points (6, 12, and 24 h) revealed a consistent difference in spontaneous migration between the two LAD-1 patients and two control subjects (data not shown).



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  		Figure 5. DC chemotaxis and transendothelial migration in LAD-1 patients. (A) CCL19/MIP-3ß (100 ng/ml) or medium alone was placed in the lower wells of a microchemotaxis chamber. DCs (5x105 cells/ml in RPMI 1640 containing 1% BSA) obtained from normal subjects or from Patients A and B were added in the upper wells. The two wells were separated by a 5-µm pore size polycarbonate filter. After incubation at 37°C in air with 5% CO2 for 120 min, filters were removed, fixed, and stained. The results are expressed as the mean (±SD) number of cells that migrated across the filter in six HPF counted in triplicate. Statistical analysis of the chemotactic response to CCL19/MIP-3ß, between LAD patients and normal donors, was performed by nonparametrical analysis test. *, Significant difference (P<0.05). (B) Spontaneous transendothelial migration of DCs. DCs (2x105 cells in 200 µl complete RPMI-1640 medium) were added to the upper well before the immersion of the trans-well. The 24-well plates were incubated for 18 h at 37°C in a 5% CO2 atmosphere. The trans-well inserts were then removed, and cells transmigrated into the well were collected by centrifugation and counted as described in Materials and Methods. The tests were run in triplicates. The results are expressed as the mean (±SD) number of cells that migrated across the trans-well filter and were counted in triplicate. DC migration among LAD patients and normal donors was compared using a nonparametrical analysis test. *, Significant difference as compared with normal controls (P<0.05).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we investigated the function and phenotype of DCs in three children with LAD-1. Analysis of lymph node structure in two of these patients revealed that factor XIIIa+ DCs, which reside in the interfollicular area of the lymph nodes, are severely reduced. Generation of DCs in vitro from monocytes obtained from LAD-1 patients confirmed that cells cultured in the presence of cytokines IL-4 and GM-CSF plus TNF-{alpha} acquired all markers of mature DCs including CD86 and CD83. These cells displayed intracellular expression of factor XIIIa, suggesting that reduction in the number of interstitial DCs in lymph nodes was not a result of altered expression of the antigen. In addition, chemotactic response to CCL19/MIP-3ß and transendothelial migration of DCs derived from LAD-1 patients were severely impaired as compared with control cells expressing normal amounts of CD18. This suggests that ß2 integrins are strictly required for migration of DCs from circulation to lymphoid tissues. Alternatively, it can be speculated that the observed decrease in the number of factor XIIIa+ DCs in the traffick area of the lymph node may be a result of a reduction in the number of recently immigrated monocytes in the lymph node—a hypothesis that could be evaluated in animal models of CD18 deficiency.

Our observation that monocytes and monocyte-derived cells are extremely sensitive to reduction in CD18 expression on cell surface suggests that the molecule is not redundant for migration of these cells from bloodstream to tissues. This finding confirms a previous report by D’Amico et al. [5 ] showing that incubation of DCs with neutralizing antibodies directed against CD18 prevented adhesion and transendothelial migration of monocyte-derived DCs. Moreover, Schneeberger et al. [6 ] have reported that CD18-/- null mice display a partial defect in accumulation of immunolabeled, interstitial DCs in the lung, indicating that adhesion and migration of monocyte-derived DCs are site-specific and depend, at least in part, on the functional integrity of CD18 molecules.

In lymph nodes from Patients B and C, a large number of LC and IDC were stained, respectively, by DC-LAMP and Langerin, in the paracortex, suggesting that DCs originated from the epidermis may use a different mechanism for migrating via lymphatic vessels. By using an assay of reverse transendothelial migration, Randolph et al. [24 ] have demonstrated that migration of skin-derived DCs via dermal lymphatic vessels does not require CD18, suggesting that the two DC subsets originated from blood or resident in tissues use distinct molecular mechanisms to migrate through various tissues.

Monocyte-derived DCs express other adhesion antigens such as DC-SIGN. However, although DCs generated from LAD-1 patients expressed DC-SIGN on the cell surface, this was not sufficient to promote adhesion and transendothelial migration of monocyte-derived DCs in the absence of CD18.

Factor XIIIa+ DCs reside in several organs and in the interfollicular area of lymph nodes where they interact with naive T cells and B cells that populate these areas, inducing Ig synthesis [14 ]. The reduced number of this DC subset may partially account for the depletion of T cells in the paracortical areas of lymph nodes and for the defective production of Ig that have been observed in LAD-1 [19 , 25 ]. As DCs are required for T cell activation during primary response to antigens, a defective recruitment of monocyte-derived DCs to lymph nodes might result in impairment of primary responses against antigens in LAD-1 patients. Other functional activities of DCs such as endocytosis of macromolecules were normal in LAD-1 patients, as immature DCs were capable of uptake and internalize FITC-dextran particles. We are investigating the antigen-presenting function of monocytes and DCs of LAD-1 subjects in mixed lymphocyte reactions with allogenic T cells; preliminary evidence indicates normal APC activities in the subjects studied. In contrast, CD18-null mice display an impairment in mixed leukocyte reaction, suggesting that functions of ß2 integrins may be different among species [26 ]. Our finding that migration of monocyte-derived DC subset cells is impaired in LAD-1 patients provides a possible explanation for the observation that development of primary immune response against novel antigens such as phage {phi}X-174 is defective in LAD-1 patients, but subsequent restimulation may result in normal activation of B and T lymphocytes [25 , 27 ]. This finding may explain the common observation that LAD-1 patients display the highest susceptibility to infections in the first years of life but improve in the following years. Investigation of primary immune response in LAD-1 subjects will be helpful to clarify this issue.


   ACKNOWLEDGEMENTS
 
This work was partially supported by the European Union Biomed-2 grant CT-98-3007, Cofinanziamento ‘98 MURST (n. 9806224439-005), CNR (PF Biotecnologief), MURST (Centro per l’Innovazione Diagnostica e Terapeutica, IDET), and ISS (grant on Secondo progetto Tuberculosi: 96/D/T27). We are grateful to Dr. Pranab K. Das for helpful discussion. The technical support of Ms. A. Galletti is also appreciated.

Received November 14, 2001; revised May 11, 2002; accepted May 21, 2002.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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