Prolactin receptor expression by lymphoid tissues in normal and immunized rats

MoEeeular and Cellular E~docri~olo~, 92 ( 1993) R21 -R25 0 1993 Elsevier Scientific Publishers Ireland, Ltd. 0303-7207/93/$06&I

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MOLCEL 02991

Rapid Paper

Prolactin receptor expression by lymphoid tissues in normal and immunized rats CfystafY. Koh a and 3. Theodore Fhilfips ‘AI

Kq W&K Prolactin receptor; Anti-receptor

antibody; Lymphocyte; Flow cytometry; (Rat)

Prolactin receptor (PRLr) expression and distribution in thymus, spleen, bone marrow, lymph nodes, and peripheral blood lymphocytes from young adult Lewis rats are analyzed using singfe-color ffow cytometry and a we&characterized monoclonal antibody directed against the rat liver PRLr. The in viva effects of regional immunization on PRLr expression are also examined. PRLr is found to be widely distributed among cells of the immune system and demonstrates Iympboid tissue-specific patterns of expression. Footpad immun~ation caused the rapid, but transient, induction of PRLr expression in the draining lymph node, with only modest effects on PRLr expression in other distant Iymphoid tissues. These studies indicate that PRL may be capable of direct interaction with the immune system through differentia1 expression of the PRL cell surface receptor on seIect iymphoid target cell populations.

Introductian Severai recent reviews have emphasized possibfe immunore~lato~ roles for proIactin (PRLj in addition to its classical endocrinologic roles (see reviews by Chikanza and Panayi, 1991; Friesen et al., 1991; Gala, 1991). Consistent with accumulating evidence that supports a direct action of PRL on various lymphoid tissues, specific PRL binding sites have been detected on human peripheral blood lymphocytes (Russell et al., lQSS> and PRL receptor (PRLr) mRNA has been recently isolated from mouse thymus and spleen (O’NeaI et af., 1991)and human T and B Iymphocytes, monocytes, and thymus ~Pell~grini et al., 1992). In this study, we have investigated the surface expression of PRLr on different Iymphoid cell types from normal, unimmunized rats and have examined possible changes in lymphoid PRLr expression following in vivo immunization using a well-characterized anti-PRLr monoclonal antibody and single-color flow hornets, We report that I~phQid cells from unimmunized rats express PRLr in a tissue-specific manner and that immunization with a

Correspondence to: J. Theodore Phiflips MD PhD, Department of Neuroiogy, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75235-9036, USA. Tel: f214) 688-4591; Fax: (214) 688-7992.

conventional, T cefl dependent antigen dramatically alters the PRLr expression pattern in the regional draining lymph node. The results establish stable, differential expression patterns of PRLr on various primary and secondary lymphoid tissues from the unimmunized adult rat, and show a dynamic response in regional iymphoid PRLr expression following in vivo immunization. Materials and methods Six to 12 week old female Lewis rats (Harlan, Indianapolis, IN) were used in al1 experiments. Rats received food and water without restriction and were acclimated to a controfled environment (12 h light; 22°C) for 1 week prior to any experiments. Operative procedures were performed under ketamine (0.2 mg/g BW s.c.> genera1 anesthesia. Experimental protocols were approved by our Institutional Review Board for Animal Research; care and maintenance were in accordance with the NIH Guide for the Care and Use of Laboratory Animals. Viable single cell suspensions of thymus, spleen, and pooled cervical, brachial, mesenteric, and popliteal lymph nodes (LN) were prepared in a standard fashion by mincing against. No. 200 wire mesh in Dulbecco’s phosphate buffered saline (Sigma, St. Louis, MO) con-

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taining 10 mg/ml bovine serum albumin (Sigma; DPBS-BSA). Parathymic LNs were removed from each thymus prior to thymus cell preparation. Bone marrow (BM) cells were obtained by aspiration from rat femurs. Peripheral blood lymphocytes (PBL) were obtained from heparinized peripheral blood, collected by cardiac puncture, and density gradient centrifugation using Histopaque 1083 (Sigma), according to the manufacturer’s specifi~tions. Contaminating red cells were removed from spleen preparations by resuspension of the washed cell pellet in 10 volumes of 0.017 M Tris containing 0.14 M NH&l, pH 7.2, for 7 min at room temperature, followed by washing in DPBS-BSA, and resuspension in ice-cold DPBS-BSA at 20 X lo6 cells/ml. The mouse monoclonal antibody mAb T6 (Okamura et al., 1989), against the purified rat liver prolactin receptor, was generously provided by Dr. Paul A. Kelly, INSERM Paris. This mAb also identifies the PRLr expressed by the rat T lympho~te line Nb2 (Okamura et al., 1989). A mouse myeloma MOPC21 IgG (Sigma) was used as an irrelevant mouse immunoglobulin control. lo6 cells/sample were pretreated with 50 pg of affinity purified sheep IgG (Pierce, Rockford, IL) in 100 ~1 for 30 min at 4°C to block immunoglobulin Fe receptors and reduce nonspecific binding, washed twice with DPBS-BSA, and blot dried. Cells were then labeled with T6 or control MOPC21 (20 pg mAb/ 10’ cells in 50 ~1) for 60 min at 4°C. Labeled and washed cells were then stained with 20 ~1 of l/50 dilution of

the fluorescent secondary antibody, FITC-conjugated sheep anti-mouse gamma F(ab’), (Sigma) in the dark for 30 min at 4”C, washed twice with DPBS-BSA, and fixed with 1.0 ml 0.1% paraformaldehyde in DPBS. All blocking, primary and secondary antibody concentrations were used at optimal concentrations determined in preliminary experiments. Stained and fixed cells were analyzed using a Becton Dickinson FACScan cell analyzer (Mountain View, CA1 and LYSIS software on a Hewlett-Packard 9000 (series 300) workstation. Dead cells were detected and gated out from analysis by concurrent propidium iodide (PI) staining (PI 50 pg/ml; 10 @/sample) of each sample. Percent PRLr + measurements are reported with control MOPC21 background staining subtracted. Assignment of cell surface PRLr expression into relative low, intermediate, and high categories is based on PRLrspecific fluorescence peak area at approximately 10, 50, and 500 arbitrary fluorescence units. Each sampling recorded at least 5000 events; calculated cell population values varied by less than 5% between experiments or among individual, identically treated animals. Histograms are expressed as number of cells (y-axis) versus fluorescence intensity (log x-axis). Anesthetized lo-12 week old rats were immunized on day 0 with an emulsion containing equal volumes of BSA (1 mg/ml in DPBS) and complete Freund’s adjuvant (Difco, Detroit, MI; containing M. tuberculosis H37Ra 10 mg/ml). Each animal received 0.1 ml of the emulsion subcutaneously into the right rear footpad for

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Fig. 1. Single-color flow cytometry of normal rat I~phoid cells. T6 and control MOPC21 histograms are represented by dark and light traces, respectively. A: thymocytes; B: splenocytes; C: bone marrow cells; 0: lymph node cells; E:peripheral blood Lymphocytes.

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Fig. 2. Single-color flow cytometry of popliteal lymph node cells from single footpad-immunized rats. T6 and control MOPC21 histograms are represented by dark and light traces, respectively. A, C, E, and G are from the draining lymph node and B, D, F and H are from the contralateral lymph node each on post-immunization days 2, 4, 6, and 8, respectively.

a total of 50 pg BSA and 500 ,ug H37Ra per animal. Rats were sacrificed by exsanguination on days 2, 4, 6, 8, and 24 following immunization for determination of PRLr expression by spleen cells and individual draining or contralateral popliteal lymph nodes. For the staining of spleen cells and cells from single popliteal lymph nodes of individual immunized rats, 1.25-2.5 X lo5 cells/sample, initial blocking with 25 pg of affinity purified sheep IgG, and 10 pg mAb were used. Results

Approximately 80% of thymus cells from unimmunized, young adult rats express homogeneous, low levels of PRLr (Fig. 1A). Thymocytes have been analyzed from female rats 2 to 6 weeks old without detectable change in degree or pattern of PRLr expression (not shown). Spleen cells demonstrate a tripartite low, in-

termediate, and high PRLr expression on approximately 35, 20, and 15% of cells, respectively (Fig. 1B). BM cells express a homogeneous level of PRLr on 50% of cells at a higher, intermediate intensity than seen on thymocytes (Fig. 10. LN cells express PRLr on 65% of cells with most expressing homogeneous low levels of PRLr and a small remainder showing an equal distribution in intermediate and high ranges (Fig. 10). PBLs also express PRLr on about 65% of cells; however, a higher proportion expresses intermediate PRLr levels compared to LN cells (Fig. 1E). We have tested several other well characterized anti-PRLr mAbs (Okamura et al., 1989) with comparable results. Preliminary analysis of additional inbred rat strains also shows similar patterns of PRLr expression. Effects of immunization on LN and spleen PRLr expression were assessed on days 2, 4, 6, 8, and 24 following a single rear footpad immunization with BSA % PRLR (+I

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Fig. 3. Changes in subpopulation PRLr expression on draining (DLNC) and contralateral (CLNC) lymph node cells (A) and splenocytes (B) on post-immunization days 0, 2, 4, 6, 8, and 24. The % PRLR (+) represent calculated percentages after the non-specific contribution of MOPC21 is subtracted.

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emulsified in complete Freund’s adjuvant. Fig. 2 compares the PRLr histogram profiles of cells from the regional draining popliteal lymph node (DLN; Fig. 2 A, C,E, G) versus cells from the contralateral popliteal lymph node from the same animal (CLN; Fig. 2B,D,F,H). As Fig. 2 shows, a major increase in PRLr expression occurs by day 6 at the site of most intense immune activation in the regional draining popliteal lymph node; PRLr expression at the distant contralatera1 lymph node is much less affected. We have also found similar results using another T-dependent antigen, myelin basic protein (not shown). Fig. 3 summarizes changes between day 0 and day 24 post-immunization in low, intermediate, and high PRLr-expressing subpopulations of DLN, CLN (Fig. 3A), and spleen (Fig. 3B) cells. Over the 24 day period, minimal overall changes are seen in PRLr expression by spleen cells, and a gradual overall decrease occurs in total PRLr+ CLN cells. A small and transient increase in intermediate and high PRLr expression and a decrease in low level PRLr expression is also seen at day 4 in spleen cells. In contrast, DLN cells expressing intermediate to high PRLr levels rapidly increase over the .first 6 days following immunization, while DLN cells expressing low levels of PRLr concurrently decrease. The subsequent fall in PRLr-intermediate and -high expressing cells by day 8 is accompanied by an increase in PRLrlow expressing cells towards the unstimulated baseline. Discussion The surface expression of PRLr on lymphoid cells of unimmunized and immunized female Lewis rats is demonstrated in this report using a mouse monoclonal antibody (T6) against the rat PRLr and single-color flow cytometry. In unimmunized animals, the differences in relative number of PRLr + cells present in thymus (80%), bone marrow (50%), spleen (70%), lymph nodes (65%), and peripheral blood lymphocytes (65%) clearly demonstrate the surface expression of PRLr in different lymphoid tissues. In addition, the degree of PRLr expression on cells from various lymphoid tissues exhibit different patterns (Fig. 1). This expression may relate, in part, to differences in cell maturity, differentiation, and microenvironment at each lymphoid tissue site. The patterns of PRLr expression and their variance among different lymphoid tissues support the hypothesis that PRL can exert a direct and physiological influence upon a significant portion of the immune apparatus. To determine a dynamic role of PRLr expression in activated lymphoid tissues, we examined PRLr expression over several days following immunization with a commonly used antigen, BSA. By immunizing rats in only one of their rear footpads, a direct comparison could be made between PRLr expression by cells ob-

tained from the activated draining popliteal lymph node versus cells from the unactivated contralateral node and spleen, all obtained from the same rat. These experiments show a distinct, but transient increase in overall PRLr expression in the activated lymph node. In addition, a dramatic shift towards intermediate and high levels of PRLr expression by individual lymph node cells, peaking at day 6, accompanies the overall increase in total PRLr+ cells. Less prominent changes in PRLr expression occurred in the contralateral node and in spleen. The changes in DLN PRLr expression could be due to the induction of increased PRLr expression by resident DLN cells or to DLN recruitment of intermediate and high PRLr expressing cells from peripheral lymphoid tissues, or both. These experiments do not directly address these possibilities. However, the transient changes in DLN PRLr expression are apparently independent of total cell number within the activated node, which remained elevated and unchanged from day 8 to day 24 (not shown). The preferential induction of increased PRLr expression in the DLN, and not elsewhere, within 48 hours of regional immunization argues strongly for an immunoregulatory role for PRL early in the normal immune response. We have recently begun two-color flow cytometry studies to define the phenotypic characteristics of T and B lymphocytes and monocyte/macrophages expressing PRLr in resting and immuneactivated lymphoid tissues. Our results are fully consistent with previous reports (Russell et al., 1985; O’Neal et al., 1991; Pelligrini et al., 1992) concerning PRLr expression in human and mouse lymphocytes. By examining individual cell surface phenotype by flow cytometry, our results also demonstrate a widespread and heterogeneous distribution of PRLr expression among constituent cells of primary and secondary lymphoid tissues in the normal, unimmunized rat. Furthermore, PRLr expression appears to be regulated dynamically within the immune apparatus as an early consequence of immune activation. Collectively, these studies help elucidate an important mechanistic connection between PRL and its reported actions on the immune system. Acknowledgements The authors wish to thank Dr. Paul A. Kelly for generously providing anti-prolactin receptor antibodies for these studies and Ms. Cheryl Beisert for assistance with preparation of the manuscript. This work was supported by NIH grant R29-DK39526. References Chikanza, I.C. and Panayi, G.S. (1991) Br. J. Rheumatol. 30,203-207. Friesen, H.G., DiMattia, G.E. and Too, C.K.L. (1991) Prog. Neuroendocrinol. Immunol. 4, l-9.

R25 Gala, R.R. (1991) Proc. Sot. Exp. Biol. Med. 198, 513-527. Okamura, H., Zachwieja, J., Raguet, S. and Kelly, P.A. (1989) Endocrinology 124, 2499-2508. O’Neal, K.D., Schwarz, L.A. and Yu-Lee, L.-y. (1991) Mol. Cell. Endocrinol. 82, 127-135.

Pellegrini, I., Lebrun, J.-J., Ali, S. and Kelly, P.A. (1992) Mol. Endocrinol. 6, 1023-1031. Russell, D.H., Kibler, R., Matrisian, L., Larson, D.F., Poulos, B. and Magun, B.E. (1985) J. Immunol. 134, 3027-3031.