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INSERM Unité 344, Molecular Endocrinology (S.J., C.K., M.-C.P.-V.), Centre d’Investigation Clinique, Hôpital Necker (J.-L.B., N.B.), and Université Paris V, CNRS URA 1461 (M.-C.G., M.D.), Hôpital Necker, Paris Cedex 15, 75730 France; Laboratory of Molecular Biology and Genetic Engineering, Université de Liège, Belgium (S.K.); Department of Medicine (Z.W.), Ludwig Maximilians Universität, Munich 80336, Germany
Address all correspondence and requests for reprints to: Dr. M.-C. Postel-Vinay, INSERM Unité 344, Faculté de Médecine Necker, 156 rue de Vaugirard, 75730 Paris Cedex 15, France. E-mail: postel-vinay{at}necker.fr
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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With the use of several biotinylated monoclonal antibodies, PRL receptors (PRLRs) have also been detected on human circulating lymphoid cells, and they appeared to be more widely distributed than GHRs (7).
Circulating GH-binding protein (GHBP) is a soluble short form of the GHR: it corresponds to the extracellular domain of the membrane-bound receptor (8). In many species, including man, GHBP is believed to be generated by proteolysis of the membrane-bound GHR (9, 10). As GHRs are most abundant in the liver, this tissue is likely to be the major source of serum GHBP. However, many tissues are able to produce GHBP and could thus contribute to the level of circulating GHBP. The level of GHBP measured in the serum is thought to reflect tissue receptor concentrations, but it remains to be clearly demonstrated in humans.
The possibility of measuring GHRs in vivo remains an important issue: until recently GHR status could only be evaluated via the GH binding protein, because of the lack of accessible cells in which GHRs can be detected. Now GHRs can be measured on peripheral blood mononuclear cells (PBMC), and several important questions need to be addressed: 1) is the GHR expression on PBMC regulated, and are the factors involved in the regulation the same as in other target cells; 2) does GHBP level reflect tissue GHR concentration. In animal models, many examples of regulation of the number of hepatic GHRs have been demonstrated (11). In humans, the level of plasma GHBP has been shown to vary in many pathophysiological situations, and age (12), nutrition (13), insulin (14), sex steroids (15), and GH itself have been shown to be important factors in the GHBP regulation (16).
In the present study, we have used flow cytofluorometry to measure GH and PRL receptors on PBMC as a function of age in normal individuals, and we have examined the relationship between plasma levels of GHBP and GHRs present in PBMC subsets.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell preparation
Human PBMC were prepared from whole blood buffy coats of donors
by centrifugation on Ficoll hypaque density gradients (Techgen
International, Les Ulis, France). The PBMC recovered from the interface
were washed twice (10 min, 400 x g, 20 C) in
HBSS and resuspended in PBS. Cells were counted and viability was
determined by trypan blue exclusion. The final PBMC concentration was
adjusted to 1 x 107 cells/ml and 100 µl of each
suspension was used for immunofluorescence staining.
Antibodies
PBMC surface markers were detected by direct labeling with
antihuman antibodies: anti-CD3 (clone UCHT 1, IgG1) as a marker of
mature T cells, anti-CD19 (clone J4–119, IgG1) as a B cell marker and
anti-CD14 (RP 052, IgG2a) as a marker of monocytes. All antibodies were
conjugated to FITC or phycoerythrin (PE) and were purchased from
Immunotech (Luminy, Marseille, France). Anti-GH receptor
mAb (10B8 mAb, mouse IgG1) was generated by immunization of mice with
recombinant nonglycosylated human GH-binding protein (17). Aggregated
human IgG, kindly given by Dr. R. Monteiro (Paris, France), was used to
block Fc receptors.
Streptavidin-PE (SAV-PE) and streptavidin-FITC (SAV-FITC) were purchased from PharMingen (Clinisciences, Paris, France). Isotype-matched antibody (PharMingen) was used as negative control.
Preparation of biotin conjugates
Anti-GH receptor mAb (10B8), biosynthetic hGH, provided by
Ares-Serono Laboratories, Inc. (Geneva, Switzerland), and
biosynthetic hPRL, provided by Dr J. Martial (Liège, Belgium),
were conjugated to biotin according to a technique previously
described, with minor modifications (18). Briefly,
biotinyl-N-hydroxysuccinimide ester (1 mg/ml;
Calbiochem, La Jolla, CA) was dissolved in
dimethylsulfoxide; 226 µl of the solution were mixed with 7 µl
carbonate buffer (1 M; pH 9.5) and 1 ml of a solution of
purified anti-GHR mAb or control antibody (1 mg/ml), hGH or hPRL
dissolved in PBS (1 mg/ml). After 2 h of incubation at room
temperature with gentle stirring, the biotin-conjugates were separated
from nonreacted biotin-7-N-hydroxysuccinamide ester by gel
filtration on a Sephadex G25 column. Positive labeling with
biotinylated conjugates was revealed with SAV-PE (Caltag). BSA was
biotinylated using the same technique.
Competition experiments
PBMC (1 x 106 cells) were incubated with
biotinylated hGH or hPRL (1 µg) for 120 min at 4 C in the presence or
absence of increasing concentrations of unlabeled hGH or hPRL (10–100
µg). Cells were then washed in staining medium, incubated with 20
µl SAV-PE and analyzed by FACSCan (Becton Dickinson and Co., Grenoble, France).
Flow cytometry analyses
Indirect labeling of cells for dual color analysis was performed
in microtiter plates by incubating first 106 freshly
isolated PBMC either with 20 µl (1 µg) biotinylated anti-GHR mAb
(10B8) for 30 min at 4 C or with biotinylated hPRL or hGH (1 µg) for
120 min at 4 C. In some experiments, cells were incubated first with 10
µl aggregated human IgG (10 mg/ml) for 20 min at 4 C, before specific
labeling, to block the Fc receptors. After washing with staining medium
(PBS plus 2% FCS and 0.01% sodium azide), cells were incubated with
10 µl SAV-PE and 10 µl FITC-conjugated specific mAb (anti-CD3,
anti-CD14, anti-CD19) or isotype matched control antibodies at the
appropriate concentrations. Cells were then washed twice and
resuspended in PBS for cytometric analysis. Controls included staining
with one reagent (FITC or SAV-PE conjugates) alone or conjugated with
biotinylated isotype matched antibody or biotinylated BSA for GHR and
PRLR respectively. Fluorescence analysis was performed on a FACScan
using Cell Quest software (Becton Dickinson and Co.). To
determine the intensity of labeling, that is the relative number of
receptors per cell, we used the mean value that represents the average
channel number of events within a cell population. Thus, results are
expressed in two ways: 1) as a percentage of cells expressing GHRs, or
GHR positive cells; or 2) as the mean value or intensity of
fluorescence that represents density of receptors per cell.
Data were collected from analysis of 10,000–30,000 viable cells. Dead cells were excluded by propidium iodide labeling. Background staining was usually less than 5%.
Assay of GH binding protein
Plasma GHBP was measured as the specific binding of
125I-hGH to GHBP, using gel filtration and HPLC, as
previously reported (12). Results are expressed as the percentage of
total radioactivity.
Statistical analyses
Results are expressed as the mean ± SD. ANOVA
and Tukey studentized range method were used for comparisons between
groups. Analysis of covariance was used to assess the effect of
cofactors, and comparison of adjusted means was tested using the
Bonferroni corrected t test. Stepwise multiple regression
was used to assess relationship between variables. The level of
significance was set at 0.05. All calculations were performed using the
standard BMDP program (19).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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. As expected, there is a significant
difference in height with age (F3,56= 2.87;
P < 0.05) and with sex (F1,56= 44.6;
P < 0.00005) (not shown). BMI also varies according to
age: F3,56= 3.42; P = 0.03, with higher
values found in the age groups 3 and 4 (>40 yr).
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): total binding corresponds to 27% of
labeled cells and nonspecific binding (in presence of 100 µg of hGH)
is 5.5% of cells. However, excess unlabeled hPRL is able to inhibit
some binding of biotinylated hGH to the cells confirming that hGH
interacts with both GHR and PRLR (Fig. 1B). Thus, to avoid
cross-reaction of hGH with PRLRs, we have used an anti-GHR monoclonal
antibody (10B8 mAb) and hPRL to measure GHRs and PRLRs, respectively.
As shown in Fig. 1C, biotinylated hPRL specifically binds to PBMC,
total binding being 25% of labeled cells, and 4% of cells
representing nonspecific binding in the experiment. We carried out
double labeling flow cytometry experiments: biotinylated 10B8 mAb and
FITC-hGH label the same cells (21%), and a few cells (5%) are labeled
only by hGH, representing binding of hGH to PRLRs (Fig. 2).
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is presented the
distribution of GHR positive cells in PBMC prepared from a young adult:
about 24% of total cells express GHR, with a low expression in T cells
(5%), whereas almost all B cells and monocytes are GHR positive.
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are presented the results of GHR
expression in PBMC and cell subsets, and of the PRLR expression in
PBMC, in the four age groups. The percentage of PBMC bearing GHRs is
significantly lower in group 2 (20–40 yr) than in groups 1 and 4 (Fig. 4), and it is also different according to
sex (F1,56 = 4.7; P < 0.05) with lower
values in female individuals (not shown). Intensity of fluorescence, as
expressed by the mean value, is comparable in the age groups (Table 2),
suggesting that the number of GHRs expressed per cell does not change
with age.
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. The proportion of B cells decreases with age.
In the three cell subsets, no significant variations with age are
detected in either the percentage of cells expressing the GHR or in the
GHR level per cell (Table 2). The distribution of GHRs in the cell
populations, shown in Fig. 3 for one adult individual, is confirmed in
subjects of the four age groups: the GHR expression is low in T cells,
is higher in B cells, and all monocytes are GHR positive. Consistently,
variance in the number of cells expressing GHRs is mainly accounted for
by GHR positive monocytes (P < 0.005) and B cells
(P < 0.005).
The percentages of PBMC expressing PRLRs and GHRs are comparable (Table 2). No differences with age are found in the percentage of PBMC
expressing PRLRs. However, the level of PRLRs expressed in PBMC is
significantly higher in group 2 than in group 4 (Table 2, and Fig. 5).
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Moreover, GHBP is higher in females than in males (Table 3). GHBP is positively correlated with
BMI (r = 0.41, P < 0.001) (not shown).
Differences of GHBP between groups remain significant after adjustment
for height and BMI.
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). This negative correlation extends to
GHR positive monocytes (P = 0.01) but not to GHR
positive B cells (P = 0.49).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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By double immunofluorescence staining, we could show that about 30% of total cells express GHRs, and that the receptor expression is low in T cells, whereas almost all monocytes and close to 90% B cells are receptor positive. A comparable distribution of GHRs has been found in human PBMC by the group of Rappaport (6) and by Badolato et al. (4), and in murine cells by our group (20). It should be pointed out that we detected a low but reproducible GHR expression in T lymphocytes in normal individuals, whereas the proportion of T cells expressing GHR varied from 2–20% in another report (6). In addition, the expression of GHRs on T cells was recently confirmed by cytofluorometry in the study of human thymocytes obtained from ten different thymuses. A proportion of 3–10% GHR positive cells was found in total cell suspensions, with higher frequencies in immature populations (21).
In this work, human PRL was used to measure the expression of PRLRs in PBMC, and the specificity of the binding of biotinylated human PRL to the cells is demonstrated. The percentages of cells expressing the GHR or the PRLR are comparable, in the range of 22–34%. A higher proportion of lymphoid cells were found to be PRLR positive in a previous study where U5, an anti-PRLR monoclonal antibody, was used (7). U5 has been prepared against the extracellular domain of the rat PRLR, and it clearly recognizes PRLRs from several species, including man (22, 23). However, in species other than rat, high concentrations of U5 must be used for detection; under such conditions, it appears that one or more proteins can be revealed with the mAb, as was shown by Western blot analysis of protein immunoprecipitated from female rat liver membranes (24).
The percentage of PBMC expressing GHRs was lower in individuals aged 20–40 yr than in younger (12–20 yr) and older (>60 yr) subjects. This unexpected result raises the question of the factor(s) regulating the expression of GHRs in man. Not much information has been presented concerning the developmental regulation of GHR in human tissues. Great variations in the number of GHRs in human liver membranes have been found, but no relationship with age or sex could be shown (25). The regulation of the plasma GH-binding protein, the soluble form of the GHR, has been more extensively studied: very low levels of GHBP have been shown in neonates (26) and during the first year of life; the highest mean value was found in young adults, aged 20–40 yr, and this result is confirmed in the present work (12). Moreover, GHBP levels are significantly higher in females than in males, in agreement with the report by Hattori et al. for individuals over 40 yr of age (27). No decrease in GHBP levels was detected in old subjects (>60 yr), contrary to what was found in another study (28); this result could be related to the good clinical, in particular nutritional, conditions of the group of elderly volunteers who were incorporated in our study.
It has been assumed that plasma GHBP levels reflect GHR tissue concentration. GHRs are abundant in the liver, which is likely to be the major source of plasma GHBP (29). Many tissues are able to produce GHBP, possibly contributing to the level of circulating GHBP. The low number of GHRs expressed on human PBMC suggests that these cells do not represent an important source of GHBP. We have found a negative correlation between plasma GHBP and the expression of GHRs in PBMC. This finding could support the idea that regulations of hepatic GHRs and blood cell GHRs are not parallel. Recent reports have shown that human peripheral blood cells are able to produce hGH (30); locally produced cytokines and GH itself could be the major regulators of GHR expression in cells of the immune system. Previous studies have shown that the level of expression of the receptors in circulating lymphocytes were not related to the serum concentration of the hormone but rather to cell activation events, both for the GHR in mice (31) and for the PRLR in man (32). Using RT-PCR, levels of GHR messenger RNA in lymphocytes from patients with acromegaly or GH deficiency were recently reported, and absence of correlation between GHR gene expression and circulating GH levels was shown (33).
In contrast with our findings, a positive correlation between GHBP levels and GHRs on B lymphocytes was found by Rappaport et al. (6). Their study, however, only included ten subjects, eight of them presenting with different pathological conditions.
Flow cytometric analysis of GHRs and PRLRs in circulating lymphoid cells is a helpful tool to evaluate receptor status in man. GHR expression is low in T cells, whereas 80–100% of B cells and monocytes are GHR positive. The inverse correlation found between plasma GHBP and expression of GHRs in PBMC suggests possible differential regulation of GHRs in lymphocytes and in other target cells such as hepatocytes and adipocytes. More studies, of various pathophysiological situations, are needed to better understand how GHR expression on peripheral monocytes is modulated.
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Acknowledgments |
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Footnotes |
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Received December 15, 1998.
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