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Endocrinology Vol. 141, No. 3 938-946
Copyright © 2000 by The Endocrine Society

ARTICLES

Identification of SP3 as a Negative Regulatory Transcription Factor in the Monocyte Expression of Growth Hormone1

Clifford R. Vines and Douglas A. Weigent

Department of Physiology and Biophysics, University of Alabama, Birmingham, Alabama 35294-0005

Address all correspondence and requests for reprints to: Dr. Douglas A. Weigent, Department of Physiology and Biophysics, University of Alabama, 1918 University Boulevard, MCLM 894, Birmingham, Alabama 35294-0005. E-mail: weigent{at}uab.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
A number of studies from different laboratories clearly show that cells of the immune system produce a GH molecule indistinguishable from that produced in the pituitary. A more recent finding from our studies suggests that monocytes use the same first exon and promoter sequence for the expression of lymphocyte GH as that reported for the expression of pituitary GH. In this report we have extended these results by determining that two members of the SP family of transcription factors, SP1 and SP3, bind to the region at -138/-133 bp containing a GGGAGG motif. Confirmation that this region of the monocyte GH promoter-bound SP1 and SP3 was accomplished using electrophoretic mobility shift assays with SP1 consensus and mutant probes as well as specific antibodies to SP1 and SP3. Selective mutation of the SP1/SP3 site increased basal transcription by 73%, indicating that this site is important in transcriptional inhibition. Overexpression of SP1 had no demonstrable effect on the GH promoter, whereas overexpression of SP3 caused inhibition of expression in P-388 monocyte cells. Cotransfection of P-388 cells with overexpression vectors for both SP1 and SP3 transcription factors also resulted in inhibition of basal expression. Transfection experiments in Drosophila SL-2 cells overexpressing SP1 and/or SP3 suggest that both factors repress the basal expression of GH promoter luciferase constructs and that the effect together was additive. Taken together, the results demonstrate that basal expression of monocyte GH may be negatively regulated by SP3.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GH IS PRIMARILY produced and secreted by somatotrophs of the anterior pituitary; however, numerous extrapituitary sites of GH synthesis have also clearly been identified. Such sites include neuronal cells within the CNS (1), endothelial cells of blood vessels (2), fibroblasts (3), epithelial cells of the mammary gland (4), thymic epithelial cells (1), and cells of the immune system, including T cells, B cells, natural killer cells, and macrophages (5). Also, some tissues, such as the placenta, synthesize variant forms of GH (hGH-V) (6). The potential role of GH in immunoregulation has been demonstrated for numerous immune functions and cell types both in vivo and in vitro (7, 8, 9, 10). Most of the studies conducted to date have examined the effect of exogenously added GH on selected immune responses, whereas the function of mononuclear cell-derived GH is less clear. Our previous experiments show that treatment of rat lymphocytes with a specific GH antisense oligodeoxynucleotide could decrease the amount of leukocyte GH synthesized and at the same time reduce lymphocyte proliferation (11). In studies with antibodies to GH, we measured a 2-fold decrease in the number of cells positive for insulin-like growth factor I (IGF-I), strongly supporting an important role for endogenously produced GH in the induction of leukocyte-derived IGF-I (12). In addition to these findings, we have also shown by dual immunofluorescence that the same spleen cells that produce GH also produce IGF-I (13). Although additional functions and mechanisms of action of immune cell derived-GH remain unknown, a recent study suggests that the effects of endogenous and exogenous GH may be similar and that they interact in an additive manner (14). Taken together, the results from a number of laboratories suggest that an intracrine/autocrine regulatory circuit may be important for the production and function of leukocyte-derived GH and IGF-I within the immune system and provide local tissue needs for these hormones independent of the pituitary whereas at the same time not disrupting the homeostasis of other organ systems.

Our original observation that rat spleen and thymus cells express detectable levels of GH message and protein (15, 16) has been confirmed in primary human peripheral blood leukocytes (17, 18, 19) as well as in a number of cell lines (20, 21, 22). The results of a number of studies employing numerous techniques, including bioassay, RIA, Western blot analysis, and the reverse hemolytic plaque assay, all support the idea that the GH protein from cells of the immune system is similar to pituitary-derived GH (15, 18, 19, 20, 21). Further, the reported sequence analysis of GH complementary DNA molecules obtained in rat and human lymphoid systems are identical to their pituitary counterparts (21, 23, 24). Currently, very little is known about the mechanisms involved in regulating the expression of the GH gene in cells of the immune system. At the cellular level, it has been demonstrated that GH-releasing hormone, TRH, somatostatin, GH, IGF-I, glucocorticoids, and mitogens modulate leukocyte GH expression (15, 18, 19, 20, 21). GH-releasing hormone stimulates, whereas IGF-I and somatostatin seem to inhibit, leukocyte GH synthesis and secretion (7, 12, 15, 25, 26). In another report exogenous GH was shown to augment endogenous GH secretion from nonstimulated and phytohemagglutinin-stimulated peripheral blood mononuclear cells (18). The studies by this latter group failed to find an effect by IGF-I on leukocyte GH (27). Both T and B cell mitogens have been found to enhance lymphocyte GH production (16).

The molecular mechanisms involved in transcriptional regulation of GH in cells of the immune system have not yet been identified. Considerable work has been done, however, in the pituitary, where both cis- and trans-specific elements have been identified to be important in the modulation of GH transcription. In the pituitary, the GH promoter binds the transcription factors nuclear factor-1, activating protein-2, upstream stimulatory factor, and the GC box-binding protein, SP1 (28). In addition to these ubiquitous transcription factors, full transcriptional activation of the GH gene in somatotropic cells requires binding of GHF-1 to confer tissue-specific expression (29). The GH promoter has two GHF-1-binding sites; binding to one of the sites replaces SP1, and binding of GHF-1 to both sites strongly activates transcription (28, 30). In the primate, SP1/SP3-binding sites as well as adjacent elements have been shown to contribute to basal and cAMP-stimulated transcriptional activation of the GH-V gene in trophoblasts (31). At the present time, nothing is known about the specific DNA elements and trans-acting factors responsible for tissue-specific expression and transcriptional activity of the GH gene in leukocytes. Our own previous studies suggest that leukocytes use the same first exon and promoter sequence for expression of GH as those previously reported for pituitary GH (32). Further, promoter deletion studies suggest the presence of both positive (299/193 bp) and negative (-193/-107 bp) regulatory elements (32); however, the specific factors involved in binding have not yet been identified. Although many of the transcription factors, such as SP1 and GHF-1, described to be involved in GH regulation in the pituitary have also been identified in cells of the immune system (24, 32, 33, 34, 35), their exact role, if any, in immune cell-derived GH synthesis has yet to be determined. Therefore, in this study we have begun to examine the molecular mechanisms that mediate basal transcription of the GH gene in the monocytic cell line P-388. We show here that both SP1 and SP3 transcription factors bind to the GH promoter, and that SP3 has a significant inhibitory action on basal expression of GH in a monocyte cell line.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture
All cell lines were cultured in RPMI medium supplemented with 10% FCS and penicillin, streptomycin, and mycostatin (100 U/ml). Cell viability was monitored by trypan blue exclusion. The mouse macrophage cell line P-388 and the Drosophila melanogaster Schneider SL2 cell line were obtained from American Type Culture Collection (Manassas, VA). In general, after electroporation cells were cultured for 24 h at 37 C before being harvested for luciferase experiments.

TA cloning
The eukaryotic TA cloning kit from Invitrogen (San Diego, CA) was used to clone amplified PCR products encoding fragments of the GH promoter. In this system, 4 µl of final PCR-amplified complementary DNA were used for ligation with 0.5 ng NotI linearized pCR II vector for TA cloning. Ligations were performed for 16 h at 15 C with 1 µl T4 DNA ligase (Life Technologies, Inc., Gaithersburg, MD). For transformation, 2 µl ligation mix were combined with 50 µl competent INV {alpha}F' Escherichia coli. After incubation, the transformation mix was spread onto Luria Bertoni agar plates containing ampicillin (50 µg/ml) and X-galactosidase (1 mg/plate). The resulting white colonies were picked and tested for the presence of PCR products in plasmids on gels after restriction enzyme analysis. All positive clones were subsequently sequenced.

Rat GH promoter/luciferase constructs
The PCR-amplified 536-bp fragment of the rat pituitary GH promoter (-523 to +13 bp) was used as a template to generate the various promoter deletion constructs in PCR reactions using specific PCR amplimer sets derived from published gene sequences (24) and Pfu (Stratagene, La Jolla, CA), a temperature-insensitive DNA polymerase that has proof-reading capabilities. PCRs were performed in a Perkin-Elmer Corp. (Foster City, CA) DNA thermal cycler. Generally, reactions were performed in a total volume of 0.1 ml containing 200 µM of each deoxy-NTP, 500 ng of each primer, 1–10 ng template DNA, and 2.5 U polymerase (Fisher Scientific, Pittsburgh, PA). The final reaction mixture was overlaid with 0.1 ml mineral oil (Perkin-Elmer Corp.) to prevent evaporation. A usual cycle consists of 1 min at 94 C (denaturation), 2 min at 58 C (annealing of primer), and 2 min at 72 C (extension). Thirty to 40 cycles (7 min total/cycle) were usually run over a 3- to 5-h period. Amplified samples were then analyzed on an agarose gel and stained with ethidium bromide to determine efficiency and size. The control template and primers derived from bacteriophage {lambda}, which define a 500-bp target, were always run to ensure the reliability of the procedure. The amplified product was subcloned into the TA vector (Invitrogen), and the entire region was sequenced. After EcoRI digestion and a subsequent fill-in reaction using the Klenow fragment of DNA polymerase I (Promega Corp., Madison, WI), this fragment was subcloned into the SmaI site of the basic pGL2-B luciferase vector Promega Corp.). The correct 5'- to 3'-orientation of the promoter fragments relative to the luciferase gene was confirmed by restriction mapping analysis and dideoxy chain termination sequencing with a synthetic oligonucleotide corresponding to the pGL2 vector sequence. Most of the sequencing reactions were performed in our laboratory with a Sequenase kit according to the vendor’s instructions (no. 70770, United States Biochemical Corp., Cleveland, OH). The sequence analysis for several constructs was verified by DNA sequence analysis in the core facility at Iowa State University (Ames, IA).

Overexpression vectors
Plasmids pCMV SP1 and pCMV SP3 (33, 34), encoding SP1 and SP3 transcription factors, respectively, were provided by Dr. Andrew Butler, University of Texas M. D. Anderson Cancer Center (Houston, TX).

Electrophoretic mobility shift assays (EMSAs)
Nuclear extracts were prepared as previously described (35) from P-388 cells. The protein concentration of the extract was determined by the Bio-Rad Laboratories, Inc. (Hercules, CA), assay with BSA as the standard. The single stranded competitor oligonucleotides were synthesized by Genosys (Woodland, TX), and the complementary sense and antisense strands were then annealed into double stranded DNAs to be used for EMSA. The GH promoter sense synthetic oligodeoxynucleotide corresponded to the region -150 to -120 bp with the sequence 5'-GACGCG-ATGTGTGGGAGGAGCTTCTAAATT. The double stranded DNA was end labeled by T4 polynucleotide kinase and [{gamma}-32P]ATP. Then, 0.2 ng probe (~2 x 105 cpm) was incubated with 10 µg nuclear extract at room temperature for 30 min in a 20-µl binding reaction that contained 20 mM HEPES (pH 7.8), 40 mm KCl, 0.5 mM dithiothreitol (DTT), 20 µg poly(dI-dC), 5 µg BSA, and 20% (vol/vol) glycerol. For competition studies, before the addition of radioactive probes, unlabeled competitors (Table 1Go) were added to the binding reactions and incubated with nuclear extract for 10 min at room temperature. Samples were then electrophoresed at 4 C on a 5% (vol/vol) polyacrylamide, nondenaturing gel in 1.0 x Tris-boric acid-electrophoresis buffer.


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  		Table 1. Oligonucleotides are listed in 5'- to 3'-orientation

 
Transfection and luciferase/ß-galactosidase assays
Rat promoter/luciferase plasmid DNA was isolated by alkaline lysis followed by polyethylene glycol precipitation (36). Cells were subdivided 3 days before transfection. After harvesting, the cell pellet was resuspended at 30 x 106 cells/ml in RPMI 1640 (no serum), 10 mM dextrose, and 0.1 mM DTT containing 20 µg luciferase construct plasmid DNA and 20 µg pON249ßGal plasmid DNA. A pulse of 400 mV and 960 µF was delivered to the cells in a 0.4-cm cuvette using the Bio-Rad Laboratories, Inc., Gene Pulser. After the pulse, the cells were maintained with growth medium. Cytoplasmic extracts were prepared 24 h after transfection; cells were washed twice with cold PBS and lysed in 0.4 ml lysis buffer [0.1 M KPO4 (pH 7.9), 0.5% Triton X-100, and 1 mM dithiothreitol] on ice for 15 min. Luciferase activities were determined as follows. A 75 x 12-mm polystyrene tube containing 100 µl cellular extract was placed in an Optocomp I luminometer (MGM Instruments, Inc., Hamden, CT), 200 µl assay buffer (100 mM Tricine, 10 mM MgSO4, 2 mM EDTA, 1 mM DTT, 2 mM ATP, and 0.1 mM luciferin) were injected, and peak luminescence was measured over a 2-sec window after a 1-sec delay. ß-Galactosidase activity was used to normalize for variations in transfection efficiency and was determined by incubating 100 µl cellular extract with 60 mM ß-mercaptoethanol and 1 mg/ml O-nitrophenyl-ß-D-galactopyranoside in 0.1 M Na2HPO4 (pH 7.3; total volume, 300 µl) at 37 C for 15 min. The reaction was stopped by the addition of 700 µl 0.1 M Na2CO3; absorbance at 410 nm was measured on a spectrophotometer. The luciferase activity of a particular construct was divided by that of the control promoterless pGL2 basic vector, and the quotient was expressed as relative luciferase activity.

Data analysis
Significant differences between various experimental treatments were determined by a test for equality of regression slopes. ANOVA and Student’s t test were used to establish the statistical significance of the results. For in vitro experiments, each fusion construct and controls were transfected at least four times, and each transfection was performed in triplicate.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
In a previous report in monocytes, by analysis of promoter deletion constructs, we identified a negative regulatory element in the promoter for GH in the region between -193 and -107 bp controlling luciferase expression (32). The decrease in luciferase expression was not observed in GH3 pituitary cells transfected with the same construct. As this region was known to be important for binding of transcription factors in pituitary GH expression, we prepared an oligonucleotide probe for EMSA covering the region -150 to -120 bp, investigated the binding of nuclear proteins to this region in the monocyte P-388 cell line, and compared the results to the GH3 pituitary cell line. The data (Fig. 1Go) show one major band, two more slowly migrating bands, and a diffuse region of binding where individual bands are not clearly discernible. In contrast, in GH3 nuclear extracts two distinctly different migrating bands, another faster migrating band, and a diffuse region similar to that observed with the P-388 nuclear extract were observed. The data suggest that different complexes are formed between the nuclear extracts from the GH3 pituitary cell line and the monocyte P-388 cell line when incubated with the oligonucleotide spanning the -150 to -120 bp region of the GH promoter. All complexes were competed by excess unlabeled -150/-120 bp oligonucleotide. Examination of the sequence of this 30-bp promoter region suggested a number of potential recognition sites for DNA-binding proteins, including those for SP1 and GHF-1. Thus, to begin to characterize the nuclear complexes formed with P-388 monocyte nuclear extracts and the -150/-120 bp probe, we tested a number of cold competitor transcription factor-binding consensus oligonucleotides (Table 1Go) rich in GC content for their ability to block the formation of bands in the gel shift assay (Fig. 2Go). The data show that the consensus oligonucleotides for nuclear factor-{kappa}B, early growth response factor, and SP1 were able to effectively compete the two slower migrating complexes, whereas IK, YY1, LyfA, Gata, Oct, and GHF-1 were essentially without effect. As the oligonucleotide probe spanning -150/-120 bp contains an SP1-like binding site (GGGAGG; GA box) at -136/-131 bp, we decided to investigate the role of this site more directly in the binding by P-388 monocyte nuclear proteins. Therefore, we prepared smaller (10-bp) oligonucleotides covering the -150/-120 bp region as well as a mutant (GGGAGG->TTTCTT) SP1 oligonucleotide and determined the effects of these unlabeled competitor oligonucleotides on complex formation with monocyte nuclear extracts (Fig. 3Go). The results of gel shift with the smaller competitor oligonucleotides show that only the -140/-130 bp region, not the -130/-120 bp region or the -150/-140 bp region, was an effective inhibitor of band formation (Fig. 3AGo). Further, the mutant SP1 30-bp oligonucleotide did not block formation of the two slower migrating bands at 10-, 20-, or 40-fold excess levels of competitor DNA (Fig. 3BGo).



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  		Figure 1. Mobility shift analysis of nuclear factors binding to the -150/-120 bp region within the GH promoter. Ten micrograms of P-388 and GH3 nuclear extracts were incubated in the absence and presence of unlabeled -150/-120 bp 10-fold excess competitor.

 


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  		Figure 2. Mobility shift assays with labeled -150/-120 bp rGH promoter oligonucleotide probe incubated with P-388 mouse monocyte cell nuclear extracts (10 µg). A 10-fold excess of each nonradiolabeled competitor (Table 1Go) was added as indicated.

 


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  		Figure 3. A, Mobility shift assays with labeled -150/-120 bp rGH promoter oligonucleotide probe incubated with P-388 mouse monocyte cell nuclear extracts (10 µg) and a 10-fold excess of 10 bp nonradiolabeled competitor oligonucleotides. B, Mobility shift assays with labeled -150/-120 bp rGH promoter oligonucleotide probe incubated with P-388 mouse monocyte cell nuclear extracts (10 µg) and 10-, 20-, and 40-fold excesses of nonradiolabeled 30-bp competitor oligonucleotides. SP1 is a consensus SP1-binding site oligonucleotide, and the SP1 mutant oligonucleotide contains a GGGAGG->TTTCTT base substitution.

 
The identities of the bands were further investigated with antisera to the zinc transcription factors SP1 and SP3, which bind SP1 recognition sites (37). Supershift analysis (Fig. 4Go) showed that the slower migrating complex was completely supershifted by antiserum to SP1. Incubation with antiserum to SP3 eliminated the lower band, and in the presence of both antisera, negligible amounts of the two original DNA-protein complexes were observed. No bands were shifted in response to antibodies to early growth response factor and YY1 (data not shown). Taken together, these data suggest that SP1 and SP3 transcription factors account for the top two slower migrating bands binding to the -150/-120 bp oligonucleotide probe.



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  		Figure 4. Mobility supershift analysis with a labeled -150/-120 bp rGH promoter oligonucleotide probe incubated with P-388 mouse monocyte cell nuclear extracts. Extracts (10 µg) were preincubated with 1 µg (1 µl) of the indicated antibody for 30 min at room temperature before the addition of radiolabeled probe.

 
Expression analysis of wild-type and mutant -150/-120 bp reporter gene plasmids in P-388 cells
To more directly examine the role of the -138/-133 bp SP1/SP3 site in GH transcription, we prepared a luciferase reporter construct with the -150/-120 bp GH promoter containing the 6-bp substitution previously shown to disrupt SP1 and SP3 binding (GGGAGG->TTTCTT). Control consensus SP1 oligonucleotide (30-mer) and the mutant SP1 oligonucleotide (30-mer) were blunt end cloned into the SmaI site of the pGL2 basic luciferase vector and along with ß-galactosidase plasmids transiently transfected by electroporation into P-388 cells. As controls, the same vector containing the full-length GH promoter and the promoterless pGL2 basic vector were also examined. Figure 5Go shows that the 6-bp substitution (GGGAGG->TTTCTT) in the consensus SP1 oligonucleotide resulted in an increase in basal transcriptional activity of approximately 73%. Although this value did not quite reach statistical significance, the trend is consistent with the ability of this site to antagonize transcription (P < 0.08). The possibility that the altered sequence in the SP1 mutant oligonucleotide had created a site that bound another factor and may have enhanced luciferase expression was also examined. Figure 6Go shows the results of an experiment with radiolabeled SP1 control and mutant oligonucleotides in gel shift after incubation with P-388 monocyte cell extracts. The data show that no new additional complex was observed with the mutant oligonucleotide in gel shift analysis and that the two slower migrating bands were not present as expected.



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  		Figure 5. Mutational analysis of the SP1/SP3 site in the GH promoter. Luciferase constructs containing GH promoter DNA were transiently transfected into P-388 cells. The constructs included either the pGH -150/-120 bp wild-type or the mutant with the GGGAGG->TTTCTT substitution at -138/-133 bp. The data are presented as a ratio to the pGL2 promoterless control. All cells were cotransfected with 25 µg/ml reporter plasmid DNA and 25 µg/ml pCMV ß-gal as an internal control for transfection efficiency. Cells were harvested as described in Materials and Methods. The results are the mean ± SE of three independent experiments. *, Significantly different from the control pGH -150/-120 construct (P < 0.08).

 


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  		Figure 6. Mobility shift analysis of nuclear factors binding to the control -150/-120 bp region and the mutant -150/-120 bp region of the GH promoter. Ten micrograms of P-388 nuclear extracts were incubated with radiolabeled probes and cold competitors as outlined in the figure and described in Materials and Methods.

 
Effect of SP1 and SP3 overexpression on GH promoter activity in P-388 cells
To further confirm the inhibitory effect of SP1 family members on GH promoter activity in P-388 cells, transient cotransfection experiments were performed with the GH promoter luciferase construct (-523/+13 bp) and SP1 and SP3 overexpression plasmids (Fig. 7Go). Overexpression of SP1 in P-388 cells had no significant effect on GH promoter activity, whereas cotransfection with SP3 significantly inhibited GH promoter activity (P < 0.05). Furthermore, cotransfection of equal concentrations of SP3 together with SP1 also resulted in an inhibition of the GH promoter by SP3. As the contributions of endogenous levels of SP1 and SP3 were unknown in these studies, we decided to study the effect of overexpression of SP1 and SP3 in the Drosophila SL2 cell lines. SL2 cells were used because they do not contain endogenous SP1 or other members of the SP family (37, 38). Surprisingly, the data (Fig. 7Go) show that both SP1 and SP3 repressed basal expression of the GH promoter. The data also show that the effects of both factors together were additive and suggest the possibility that SP1 may interact with another tissue-specific transcription factor in P-388 cells that modulates its inhibitory effect. A control experiment in the SL2 cell line was also performed, examining the effect of overexpression of GHF and Oct proteins on GH promoter luciferase expression to rule out a nonspecific inhibitory effect in the SL2 cell line. The data showed that overexpression of either of these two transcription factors was similar to that of the control SP vector alone, in that neither factor had any effect on luciferase expression (data not shown).



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  		Figure 7. Inhibition of GH promoter luciferase activity by SP3 in P-388 and SL2 cells. P-388 cells were electroporated with 25 µg/ml GH reporter plasmid (pGH -523 +13) and 50 µg/ml pCon DNA, 25 µg/ml pCMV-SP1 or pCMV SP3, or 25 µg/ml of both pCMV-SP1 and pCMV-SP3 and 25 µg/ml pCMV ß-galactosidase as an internal control for transfection efficiency. The total amount of transfected DNA was kept constant by adding control plasmid (pCon DNA). Cells were harvested as described in Materials and Methods. The results are the mean ± SE of three independent experiments. *, Significantly different from the control nonoverexpressing group of transfected cells (P < 0.05).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The expression of GH, once thought to be exclusive to the pituitary somatotrope, has now been extended to include cells of the immune system. The data, using numerous techniques, including bioassay, RIA, Western blot analysis, and the reverse hemolytic plaque assay, show that the GH protein from lymphocytes is similar to pituitary-derived GH (7). Our data suggest that the same transcription initiation site is used in both pituitary and mononuclear cells; therefore, both cell types appear to use the same 5'-flanking region to govern the expression of the GH gene (39). The same promoter was shown to be used in the case of hypothalamic and lymphocyte expression of LHRH (40) and pituitary and monocyte expression of POMC (41). Alternative promoters, however, have been described for human PRL and rat GH-releasing hormone expression outside the neuroendocrine system (42, 43). In another study, pertinent to the results presented here, the Northern analysis of messenger RNA (mRNA) in bovine lymphoid cells indicated that lymphoid GH mRNA was approximately 350 nucleotides larger than that in the pituitary (44). The coding regions and 3'-untranslated regions of the bovine lymphocyte GH and pituitary transcripts were the same; however, analysis of the 5'-untranslated region showed that the transcription began upstream from the start site in the pituitary gland (44), The reason for the difference between the bovine and rat data are not clear; however, it may stem from the fact that our work was performed in a mouse monocyte cell line, whereas the bovine findings were obtained in fetal lymphoid cells and adult peripheral blood lymphocytes.

The GH, GH-V, and CS genes contain SP1-binding sites approximately 136 bp from the start site of transcription, and the importance of SP1/SP3 binding elements for activation of the GH gene has been previously demonstrated (31). In the monkey, mutagenesis studies show that the distal and proximal SP1/SP3 binding elements contribute to transcriptional activation of the placental monkey GH-V gene. In this same study, elements adjacent to the SP1/SP3 sites were shown to be critical for both basal and cAMP-stimulated transcription in placental cells (31). A previous cell-free transcription study has shown that SP1 can displace GHF-1 from its distal binding site and stimulate transcription from the human GH gene promoter (30). The interplay between SP1 and GHF-1 has been interpreted as a fine-tuning mechanism by which at low GHF-1 concentrations, SP1 is able to bind to the promoter and in part compensate for the decreased stimulation by GHF-1 (30). Here we have demonstrated that both SP1 and SP3 bind to sites in the monocyte GH promoter and form two complexes by EMSA. The slower moving band was recognized and supershifted by an antibody specific to SP1. The faster moving band results from binding of SP3, as antibody to SP3 interfered with the formation of this complex. Both complexes were supershifted when antisera to SP1 and SP3 were used together. Consistent with the two proteins acting at or near the same site was our ability to block the formation of both complexes with a control oligodeoxynucleotide to the SP1 site, but not a mutant oligodeoxynucleotide.

It is generally accepted that SP1 stimulates transcription, and SP3 represses SP1-mediated transcriptional activation, suggesting that SP3 is an inhibitory member of the SP family of transcription factors (45). Our studies in Schneider’s Drosophila SL2 cells, in contrast to findings for P-388 cells, demonstrated that both SP1 and SP3 inhibit the GH promoter. These data suggest that the levels of SP1 and SP3 or the SP1/SP3 ratio may play a role in regulating the transcriptional activity of the GH promoter in monocytes. The discrepancy between the data obtained in the SL2 cell line and P-388 may be explained by the differing endogenous levels of SP1 in the different cell types or the different levels of SP1 and SP3 proteins produced by each particular overexpression plasmid. The overexpression of SP1 and SP3 proteins was demonstrated by immunoblot analysis of nuclear extracts (data not shown), although the exact concentrations have not been determined. It has also recently been shown that SP3 encodes three distinct proteins in vivo that differ in their capacity to stimulate or repress transcription (46). Thus, the levels of SP1 as well as SP3 isoform will need to be determined to assess whether this may explain the difference in the two systems.

It is becoming clear that SP1 binding and trans-activation are regulated by several stimuli that are important in the regulation of cellular growth and function (47). The SP1 transcription factor is found in glycosylated and phosphorylated forms, but little is known about how these modifications affect function (48, 49). Also, interactions between the retinoblastoma (Rb) protein and SP1 have been reported (50) as well as interactions of SP1 with Oct 1 (51). Other factors with which SP1 is known to interact include SF-1, p53, STAT1 (signal transducer and activator of transcription-1), Gata-1, activating protein-1, nuclear factor-{kappa}B, and the estrogen receptor (52, 53, 54, 55, 56, 57). These reports suggest that there may be multiple mechanisms in cells that influence the interaction of SP1 with the transcription factor IID complex (58). It seems the data from a number of different systems demonstrate that depending upon the context and/or the number of functional G/C boxes present, cell cycle-regulated promoters display a selective responsiveness to the SP1/SP3 ratio (47). In a preliminary study we observed no effect of overexpression of Rb protein in P-388 cells on GH promoter activity (unpublished). Overexpression of GHF and Oct proteins in P-388 cells, however, resulted in a modest inhibition of GH promoter activity (32). Clearly, further investigations will be required to unravel the mechanisms and interplay between cis-elements and trans-acting factors controlling the gene expression of monocyte GH and the role this hormone plays in the immune response. Nevertheless, it is tempting to speculate that this hormone in monocytes may play a role in cell-cycle progression.

The region of the rat GH (rGH) 5'-flanking sequences contained between positions -237 and -47 bp direct tissue-specific expression of GH in the pituitary (29). This region of the rGH gene contains two binding sites for the pituitary-specific factor Pit-1 as well as sites for other factors (59). The pituitary homeodomain transcription factor Pit-1 serves an important role in the trans-activation of the GH gene in the pituitary (29). It has also been shown that Pit-1 is expressed in hemopoietic and lymphoid tissues (60). However, the idea that Pit-1 may not be involved in GH expression in the murine system was first suggested by our work showing near-normal levels of GH mRNA and protein in dwarf spleen cells compared with those in the pituitary in these animals (61). This work was essentially confirmed and extended to bone marrow cells, where in situ hybridization, immunocytochemistry, and RT-PCR analysis showed that GH expression does not depend on Pit-1 (62). In preliminary experiments using EMSA we have not been able to shift any band formed by complexes with proteins from P-388 nuclear extracts and GHF oligonucleotides or GHF-specific antibodies (data not shown). A similar situation has been described for human and monkey trophoblasts, in that although GHF-1 expression can be detected, supershift analysis could not detect GHF-1 binding to this region (31). It may be that under certain conditions, selected cells of the immune system may use Pit-1 to regulate GH, whereas it does not appear to be required for basal expression.

There exists a considerable body of literature that attests to the interaction between GH and the immune system (7). Animals with a genetic defect leading to decreased secretion of pituitary GH and other pituitary hormones are stunted in growth and have an immune deficiency (63). Increased growth of the spleen has been observed by treating normal rats, hypophysectomized rats, and Snell dwarf mice with GH as well as in transgenic mice overexpressing GH (64, 65). Although GH corrected some of the hemopoietic deficiencies in dwarf mice (66), it failed to restore the frequency of B cell progenitor populations to normal, which required T4 (67). Also, GH influences granulopoiesis, erythropoiesis, and immune function in vitro and in vivo in animal models (63). Despite this, however, GH hypo- or hypersecretion in humans does not induce clinically significant immunological impairment (68). Subclinical changes in chemotaxis, phagocytosis, and natural killer cell activity, however, have been reported in children with GH deficiency (69). The normal immunocompetence of humans with GH deficiency can in part be explained by the presence of normal serum PRL levels as well as the presence of interleukin-2 and interferon-{gamma}, which mediate similar and redundant effects in the immune system. Although not much information is available on the immunocompetent state of the Laron dwarf, the clinical profile is indistinguishable from that of isolated GH deficiency (70). There is evidence in Laron T cell lines that lack a functional GH receptor for an alternative pathway of GH action through intact PRL receptors (71, 72). Thus augmentation of colony formation and insulin resistance induced by GH in Laron dwarf T cells may occur via lactogenic receptors in the absence of the GH receptor.

The expression of GH and its receptor on lymphoid cells suggests that GH exerts physiological effects on the immune system. Most of the studies conducted to date, however, have examined the effect of exogenously added GH, whereas the function of endogenous lymphocyte-derived GH on selected immune responses is less clear. Our previous studies showed that a specific GH antisense oligodeoxynucleotide could decrease the amount of leukocyte GH synthesized and lymphocyte proliferation (11). In blocking studies with antibodies to GH we observed a 2-fold decrease in the number of cells positive for IGF-I, strongly supporting an important role for endogenously produced GH in the induction of leukocyte-derived IGF-I. We have also shown that the same cells that produce GH also produce IGF-I (13). The idea that GH produced by lymphocytes may function primarily in a paracrine or autocrine, rather than endocrine, role is supported by the low levels of GH produced by immune cells. In the case of GH, it appears that approximately 2% of lymphoid cells are positive by immunofluorescence, whereas only 0.1% of cells are secreting GH, as determined by the reverse hemolytic plaque assay (20). Thus, much more of the lymphocyte hormone appears to remain in the cell and may function in an intracellular fashion, like PRL, or in an autocrine manner (12, 73). Taken together, the results suggest that an intracrine/autocrine regulatory circuit may be important for the production of leukocyte-derived GH and IGF-I within the immune system and provide local tissue needs for these hormones independent of the pituitary.


   Acknowledgments
 
We thank Diane Weigent for excellent editorial assistance and typing the manuscript.


   Footnotes
 
1 This work was supported in large part by grants from the National Institute of Neurology and Communicative Disorders (RO1-NS-24636) and the NIDDK (RO1-DK-38024). Back

Received March 25, 1999.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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