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Department of Physiology, Goteborg University (C.L., M.G., K.D., H.W.), SE 405 30 Goteborg; Research Center for Endocrinology and Metabolism, Department of Internal Medicine, Sahlgrenska University Hospital (G.H., B.C.), SE 413 45 Goteborg; and Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Goteborg University (H.B.), SE 405 30 Goteborg, Sweden
Address all correspondence and requests for reprints to: Dr. Håkan Billig, Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Goteborg University, P.O. Box 434, SE 405 30 Goteborg, Sweden. E-mail: hakan.billig{at}obgyn.gu.se
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The PRL receptor (PRLR) belongs to the cytokine receptor superfamily and is widely distributed in vertebrates (1). However, with the exception of birds (1, 12), the receptor has not been detected in white adipose tissues. In other tissues different PRLR splice forms have been detected in several species (1). For instance, in the mouse, one long (L-PRLR) and three short receptor forms (S1-PRLR, S2-PRLR, and S3-PRLR) have been identified (13). Both the long and short receptor forms have been shown to dimerize and activate JAK2, Fyn, and the mitogen-activated protein kinase systems (14, 15, 16). However, only the long PRLR splice form is able to activate the Jak2/Stat5 pathway and initiate milk protein gene transcription (17). The expression of the various PRLR splice forms has been reported to be regulated differently in several tissues during the estrous cycle, pregnancy, and lactation (13, 18, 19, 20, 21, 22, 23).
Several studies indicate that PRL may be an important regulator of adipocyte metabolism. Therefore, the aim of this study was to examine the expression and possible regulation of the PRLR in adipose tissue, using the mouse as an experimental animal model.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Protein and RNA extraction
Two different protein extraction methods were used, one for
adipose tissues and another for liver tissues. Adipose tissue was
homogenized in lysis buffer [1 mM EDTA, 10 mM
Tris (pH 7.5), and 0.25 M sucrose] containing 0.6
mM phenylmethylsulfonylfluoride, 1 mM
Na2VO3, and 0.27 trypsin
inhibitor units aprotinin/ml (Sigma, St. Louis, MO) and
centrifuged (15,000 x g, 10 min, 4 C), the interphase
was saved (the upper phase contains triglycerides and FFA), 1%
Triton X-100 was added, and the samples were put on a rocking cradle
for 1 h at 4 C and then centrifuged (15,000x g, 50
min, 4 C). The supernatant was stored at -70 C until analyzed. Liver
tissues were homogenized in lysis buffer (25 mM
Tris-HCl and 0.3 M sucrose, pH 7.4) containing
200 kallikrein inhibitory U/ml aprotinin, 1 mg/ml leupeptin, 1 mg/ml
pepstatin, and 1 mg/ml Pefablock (Roche Molecular Biochemicals, Mannheim, Germany) and centrifuged (800 x
g, 20 min, 4 C), and the supernatant was saved and
centrifuged (100,000x g, 60 min, 4 C). The pellet was
resuspended in the lysis buffer together with 1% Triton X-100, put on
a rocking cradle for 1 h at 4 C, and then centrifuged (15,000x
g, 60 min, 4 C). The supernatants were stored at -70 C
until analyzed. The protein content in all samples was measured using
the Bio-Rad Laboratories, Inc., protein assay (Life Technologies, Inc., Gaithersburg, MD).
Total RNA was extracted from frozen tissues by acid guanidinium thiocyanate-phenol-chloroform extraction as described by Chomczynski and Sacchi (26).
Preparation of isolated adipocytes
Adipocytes were isolated from adipose tissue by collagenase
treatment (Sigma) (27). Adipocytes were
filtered through a nylon mesh (250 µm pore size) to remove undigested
tissue fragments and stroma (the stromal fraction). The adipocytes,
which float to the surface of the medium, were washed three times and
studied under a microscope, making sure they were adipocytes. RNA was
extracted from the adipose tissue, the adipocytes, and the stromal
fraction, respectively.
RT-PCR
Complementary DNA (cDNA) was synthesized using 1 µg total RNA
together with 0.25 µg oligo(deoxythymidine) primer (0.5 µg/µl),
in a total volume of 5 µl, which was denatured at 70 C for 5 min.
After annealing at room temperature for 10 min, 15 µl RT mix
containing 20 U RNasin (Promega Corp., Madison, WI), 5 U
AMV reverse transcriptase (Promega Corp.), RT buffer
(Promega Corp.), and 20 mmol deoxy-NTP (Roche Molecular Biochemicals) were added. cDNA synthesis was carried
out at 42 C for 60 min.
The expression of the 4 different PRLR splice forms L, S1, S2, and S3,
was analyzed using 1 common PCR primer, prlr1, from the extracellular
domain (nucleotides 610–632; 5'-AAGCCAGACCATGGATACTGGAG-3'; position
is shown in Fig. 1
) (13),
and 4 specific primers, pL, pS1, pS2, and pS3, corresponding to the
first 23 nucleotides of the unique coding region of the different
receptor splice forms (nucleotides 841–863; pL,
5'-AGCAGTTCTTCAGACTTGCCCTT-3'; pS1, 5'-AACTGGAGAATAGAACACCAGAG-3'; pS2,
5'-TCAAGTTGCTCTTTGTTGT GAAC-3'; pS3, 5'-TTGTATTTGCTTGGAGAGCCAGT-3';
positions are shown in Fig. 1) (13) in 4 different PCR
reactions/tissue, generating 254-bp PRLR-PCR products, together with
PCR primers for the internal standard the ribosomal protein L19
(L19-fw, 5'-TGCGGGAAAAAGAAGGTCTGGTT-3'; L19-rev, 5'GCG
AGCCTCAGCCTGGTCAG-3') (28), generating a 438-bp L19-PCR
product. The following PCR reactions were optimal for liver and adipose
tissue, respectively; 94 C for 2 min, then 26 or 29 cycles of
sequential incubations at 94 C for 45 sec, 59 C for 1 min, 72 C for 1
min, and finally extension at 72 C for 7 min. The PCR products were
separated on a 2% 0.5 x TBE [1 mM EDTA
(pH 8) and 45 mM Tris-borate]-agarose gel together
with a 100-bp ladder (Promega Corp.). The different PRLR
PCR products were verified. They were cloned into the pCRII-TOPO vector
(Invitrogen, Carlsbad, CA) and sequenced.
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RNA probes
A 288-bp mouse PRLR cDNA fragment
(HincII-BamHI, nucleotides 1507–1794)
(13) subcloned into a pBluescript vector was used as a
template. This PRLR cDNA fragment is part of the intracellular PRLR
cDNA and will only detect L-PRLR when used as a probe in the
ribonuclease protection assay (RPA) (13). The probe
position is shown in Fig. 1
. The construct was linearized using
XhoI, and an antisense
[32P]CTP-labeled PRLR RNA probe was synthesized
using T3 RNA polymerase according to the manufacturer’s instructions
(Ambion, Inc., Austin, TX).
A mouse cyclophilin template (no. 7675, Ambion, Inc.), generating a 103-bp protected fragment (nucleotides 38–140), was used as an internal standard to control the amount of RNA in each sample. An antisense [32P]CTP-labeled cyclophilin RNA probe was synthesized using T3 RNA polymerase according to the manufacturer’s instructions (Ambion, Inc.). Compared with the PRLR probe, 6.25 times less [32P]CTP was used when synthesizing the cyclophilin probe, making the cyclophilin probe less radiolabeled.
A 412-bp mouse leptin cDNA fragment (nucleotides 64–475) subcloned into a pCRII-Topo vector (Invitrogen) was used as a template. The construct was linearized using HindIII, and an antisense [32P]CTPlabeled leptin RNA probe was synthesized using T7 RNA polymerase according to the manufacturer’s instructions (Ambion, Inc.). Before use, all RNA probes were gel purified, run on a denaturing 8 mmol/liter urea-6% polyacrylamide gel (Novex, San Diego, CA), identified, excised, and eluted, and the specific activity was determined in a scintillation counter.
RPA
RPA was performed using the RPAIII ribonuclease protection assay
kit (Ambion, Inc.). Before running the samples, the PRLR
signal, the cyclophilin signal, the leptin signal, and the
PRLR/cyclophilin and leptin/cyclophilin ratios were confirmed to
increase linearly with increasing amounts of RNA (10–40 µg). When
analyzing L-PRLR messenger RNA (mRNA) expression, each sample (20 µg
total RNA) was hybridized overnight at 42 C with 140,000 cpm antisense
[32P]CTP-labeled PRLR RNA probe and 100,000 cpm
antisense [32P]CTP-labeled cyclophilin RNA
probe. When analyzing leptin mRNA expression, each sample (20 µg
total RNA) was hybridized overnight at 42 C with 200,000 cpm antisense
[32P]CTP-labeled leptin RNA probe and 100,000
cpm antisense [32P]CTP-labeled cyclophilin RNA
probe. The protected fragments were separated on a denaturing 8
mmol/liter urea-6% polyacrylamide gel (Novex) for 1
h at 130 V. After drying the gel, it was exposed to a PhosphorImager
screen for 2 days and developed in a PhosphorImager (Molecular Dynamics, Inc., Sunnyvale, CA), and quantitative analysis was
performed using ImageQuant software (Molecular Dynamics, Inc.). The
PRLR/cyclophilin ratio was calculated for each sample, and the
leptin/cyclophilin ratio was calculated in samples when analyzing
leptin expression.
Immunoblotting
Dynabeads (antimouse 112.01, Dynal, Oslo, Norway)
were coated with a specific antibody against the PRLR, AbU5 [MA1–610,
monoclonal (mouse) anti-PRLR antibody (IgG1), clone U5, Affinity BioReagents, Inc., Golden, CO], following the manufacturer’s
instructions (12.5 µg antibody/250 µl beads). PRLR protein was
immunoprecipitated by adding 60 µl AbU5-coated Dynabeads to 1 mg
protein sample and incubated on a rocking cradle at 4 C overnight, then
the Dynabeads were washed four times before the precipitated proteins
were eluted in 20 µl 1x SDS loading buffer at 100 C for 5 min. The
AbU5-immunoprecipitated samples were loaded on a SDS-polyacrylamide gel
(6% Tris-glycine gel, Novex, San Diego, CA). Prestained
standards (SeeBlue and MultiMark, Novex) were used as
weight markers. The proteins were transferred onto a polyvinylidene
difluoride membrane (Millipore Corp., Bedford, MA) using a
blotting system (Hoefer TE-series, Pharmacia Biotec, Uppsala, Sweden). The membrane was washed in WBI (twice,
15 min each time; 1 g MgCl2, 0.2 g
sodium azide, and 3 ml Tween-20 added to 1 liter PBS buffer) and
blocked in WBII (2 h; 1 g MgCl2, 0.2 g
sodium azide, 3 ml Tween-20, and 2 g I-Block (Tropix,
Bedford, MA) added to 1 liter PBS buffer, pH 7.4) with antimouse IgG
added (dilution, 1:30 000; B0529, Sigma). The membrane was
then incubated with a specific antibody against the PRLR, AbU5 (4 C,
overnight; dilution, 1:1000; MA1–610, Affinity BioReagents, Inc.), washed in WBI and WBII, and incubated with antimouse IgG
linked to alkaline phosphatase (2 h; dilution, 1:30,000; A-1682,
Sigma). Immunoreactive protein was visualized by
chemiluminescence using CDP-Star as substrate (Tropix). The membrane
was exposed to ECL film (Amersham Pharmacia Biotech) at
room temperature for 10–90 sec and subsequently developed in a Curix
60 developing machine (Agfa-Geveart AG, München,
Germany).
Statistical analysis and scanning
Differences in PRLR mRNA expression (comparing PRLR/cyclophilin
ratios) and differences in adipose tissue weight among groups were
analyzed using one-way ANOVA followed by Student-Newman-Keuls multiple
range test. When appropriate, values were transformed to
logarithms.
Polaroid photographs and autoradiograms were scanned using a Fluor-S Multi Imager (Life Technologies, Inc.).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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together with a primer pair detecting the
internal standard L19 (28). In adipose tissue of virgin
female mice L-, S2-, and S3-PRLR were detected (Fig. 2A). S1-PRLR was below the detection
limit (Fig. 2A). The liver of virgin female mice was used as a control
tissue in the RT-PCR, and all four mRNAs, L-, S1-, S2-, and S3-PRLR,
were detected (Fig. 2A). The relative abundance of the four PRLR mRNAs
detected in the liver is in agreement with previously published data
(13). Furthermore, in adipose tissue of male, pregnant,
and lactating mice, L-, S2-, and S3-PRLR were detected using RT-PCT
(data not shown). The identities of the PCR products were confirmed by
DNA sequencing. In the next series of experiments, we analyzed the
level of L-PRLR mRNA expression in adipose tissue by RPA, using a
288-bp PRLR probe (Fig. 1). A 103-bp cyclophilin probe was used as an
internal standard, which controls the amount of RNA in each sample by
calculating the PRLR/cyclophilin ratio. L-PRLR mRNA expression was
detected in adipose tissue by RPA. However, in the liver the level of
L-PRLR mRNA expression was severalfold higher compared with that in
adipose tissue (Fig. 2B).
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, +/+ Adip.tissue and
+/+Liver). The 95,000 Mr protein band could not
be detected in adipose tissue and liver extracts from PRLR
gene-deficient mice (Fig. 2C, -/-Adip.tissue and -/-Liver). L-PRLR
is believed to be 95,000 Mr (30).
The IgGH band (Mr 50,000)
is the result of the AbU5 antibody used for immunoprecipitation and
immunoblotting (30).
Cellular distribution of PRLR mRNA expression in mouse adipose
tissue
The cellular distribution of PRLR in adipose tissue was analyzed
after isolating adipocytes from the stroma by collagenase treatment as
previously described (27). Using RPA, L-PRLR expression
was detected in the isolated adipocytes, in the fraction containing
stroma and undigested tissue fragments (the stromal fraction), and in
the adipose tissue (Fig. 3). To
demonstrate the efficiency of the cell separation procedure, a protein
expressed in adipocytes, leptin, was also analyzed in the three
different samples: adipose tissue, isolated adipocytes, and stromal
fraction. The level of leptin expression relative to the expression of
the internal standard, cyclophilin, was almost doubled in the isolated
adipocytes compared with the level in adipose tissue.
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).
The pituitary gland and mammary gland were used as control tissues in
the RT-PCR. A 663-bp PCR fragment was generated in pituitary gland and
mammary gland, confirming the PRL production in these tissues. However,
no PCR fragments were generated in adipose tissue and liver,
demonstrating that there is no local PRL production in mouse adipose
tissue or liver.
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). In adipose
tissue, there was no significant change in L-PRLR mRNA expression
during pregnancy compared with that in virgin female mice. However,
during lactation the level of L-PRLR mRNA expression increased 2.3-fold
compared with that in virgin mice. Furthermore, in adipose tissue the
levels of L-PRLR mRNA expression in virgin female and male mice were
similar (Fig. 5, A and B). The regulation of PRLR mRNA in adipose
tissue was compared with its regulation in liver. In liver, a different
PRLR mRNA expression pattern was found. L-PRLR expression increased
3.4-fold during pregnancy compared with that in virgin females.
However, during lactation the L-PRLR expression level was similar to
the level detected in virgin females. A 4.5-fold higher level of L-PRLR
expression was found in the liver of virgin female mice compared with
that in male mice (Fig. 5, C and D).
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and 7).
The level of L-PRLR expression was increased 1.4-fold in adipose tissue
of female PRL-transgenic mice compared with the control value (Fig. 6, A and B). Furthermore, in the liver the level of L-PRLR expression was
increased 1.9-fold in the female PRL-transgenics compared with that in
the controls (Fig. 6, C and D). In adipose tissue of male
PRL-transgenic mice, the level of L-PRLR expression was increased
2.4-fold compared with the control value (Fig. 7, A and B).
Furthermore, in the liver the level of L-PRLR expression was increased
2.7-fold in the male PRL-transgenic mice compared with that in the
controls (Fig. 7, C and D).
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). In addition, the masses of
parametrial and retroperitoneal adipose tissues in female
PRL-transgenic and control mice were recorded. In female PRL-transgenic
mice, retroperitoneal adipose tissue was decreased in weight compared
with that in controls (Table 2). However,
there was no significant difference between PRL-transgenic and control
mice when comparing the mass of parametrial adipose tissues.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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We examined the expression and regulation of the PRLR mRNA using RT-PCR and RPA. L-PRLR is the only splice form shown to activate the Jak2/Stat5 pathway and initiate milk protein gene transcription (17). L-, S2-, and S3-PRLR mRNA expressions were detected in adipose tissue. However, L-PRLR seems to be the most abundant form in adipose tissue. Immunoblot analysis with a monoclonal anti-PRLR antibody (AbU5) (31, 32) was performed to examine PRLR protein expression in mouse adipose tissue. The long PRLR isoform has previously been shown to be 95,000 Mr (30, 33). A protein band of 95,000 Mr was detected in both adipose tissue and liver isolated from wild-type mice. The 95,000 Mr protein band was absent in both adipose tissue and liver from PRLR gene-deficient mice.
A role for the PRLR in adipocytes has been suggested in the following study. When bone marrow stromal cells were treated with adipogenic agonists, PRLR mRNA expression was induced after adipocyte differentiation, (34). Furthermore, PRL has been reported to affect LPL activity in differentiating 3T3-L1 adipocytes (35). To determine the cellular distribution of PRLR expression in adipose tissue, adipocytes were separated from the stromal fraction by collagenase treatment (27). RNA was extracted from three samples: adipose tissue, isolated adipocytes, and stromal fraction. In previous studies leptin was demonstrated to be expressed in adipocytes (36, 37). The increased level of leptin expression, which almost doubled in the adipocyte fraction compared with that in adipose tissue, indicates a large enrichment of adipocytes when isolating them using the described method (27). A very low level of leptin expression was detected in the stromal fraction, indicating a low amount of remaining adipocytes in the stromal fraction. L-PRLR mRNA was expressed at similar levels in adipose tissue, isolated adipocytes, and stromal fractions. Taken together with the expression of leptin in the cell fractions, these results suggest that PRLR is expressed in both adipocytes and stroma.
In addition, L-PRLR regulation in adipose tissue of wild-type male, virgin female, pregnant, and lactating mice was analyzed by RPA. In adipose tissue, L-PRLR mRNA expression increased 2.3-fold during lactation compared with that in virgin mice. This finding suggests a direct physiological role for PRL, mediated via its receptor, in adipose tissue. For instance, the parametrial and retroperitoneal adipose tissues decreased in weight during lactation. Therefore, the increased L-PRLR expression found in mouse adipose tissue during lactation might be involved in regulating the LPL activity and metabolic changes found in adipose tissue of rodents during lactation (7, 8, 10). However, some studies suggest that the metabolic changes found in adipose tissue during lactation are indirect PRL effects and do not require functional PRLRs in the adipose tissue (6, 38). Insulin has been reported to be an important regulator of the different metabolic changes found during lactation (6, 10, 38), and PRL might be involved in regulating the sensitivity of the insulin receptor in adipose tissue during lactation (6, 10). Furthermore, PRL has been demonstrated to affect glucose transport (3, 4) and insulin binding (3) in adipocytes cultured in vitro, and these effects are probably mediated by functional PRLRs.
The level of PRLR expression is regulated differently in several tissues during pregnancy and lactation (13, 18, 19, 20, 22, 23). To compare the regulation of L-PRLR in different mouse tissues, we also analyzed the regulation of L-PRLR mRNA in the liver of wild-type male, virgin female, pregnant, and lactating mice. In the mouse liver the level of L-PRLR mRNA increased dramatically during pregnancy compared with those in virgin and lactating mice, showing a different PRLR regulation in the liver compared with that in adipose tissue during pregnancy and lactation. In the rat liver, the level of PRLR expression has been found to increase during pregnancy compared with those in virgin and lactating rats. The opposite was detected in the mammary gland, where the level of PRLR expression was low in virgin and pregnant rats and then increased during lactation (22). There is also reason to believe that the PRLR is transcriptionally regulated, because the pattern of antibody binding to rat PRLR protein was roughly parallel to that of rat PRLR mRNA expression (22). Furthermore, one group has reported the transcripts corresponding to the short PRLR isoforms to increase during pregnancy in the mouse liver, using Northern blot analysis (18). We also found a sex difference in the level of PRLR mRNA expression. In the mouse liver PRLR expression was higher in females compared with in males. In contrast, in adipose tissue no difference in the level of L-PRLR expression was detected in female and male mice.
The regulation of PRLR is under hormonal control. For instance, estrogen and progesterone have been reported to regulate PRLR expression in liver and breast (19, 39, 40, 41). Furthermore, GH also regulates PRLR expression in mouse liver (42, 43), and T4 together with propylthiouracil have been reported to regulate the level of PRLR in female rat kidney (44). The ligand PRL is also an important regulator of PRLR expression (39). We examined L-PRLR mRNA expression in adipose tissue and liver of female and male PRL-transgenic mice and controls. In adipose tissue, the level of L-PRLR expression was 1.4-fold higher in female PRL-transgenics than in controls. However, the increase in L-PRLR expression in adipose tissue was greater during lactation (2.3-fold) compared with that in female PRL-transgenics (1.4-fold). These results suggest that PRL is an important regulator of L-PRLR expression in female adipose tissue, but the results also indicate that other regulators are of importance. Furthermore, in female PRL-transgenic mice the level of L-PRLR expression increased in the liver (1.9-fold) compared with that in controls. However, in the liver of wild-type lactating mice L-PRLR expression decreased compared with that during pregnancy. This result demonstrates that several factors are important for regulation of the level of PRLR expression in the female liver. In male PRL-transgenic mice the level of L-PRLR expression increased in both adipose tissue (2.4-fold) and liver (2.7-fold) compared with the control value. The increased L-PRLR expression in both adipose tissue (2.4-fold) and liver (2.7-fold) of male PRL-transgenics was higher than the increased L-PRLR expression in adipose tissue (1.4-fold) and liver (1.9-fold) of female PRL-transgenics. These results indicate a stronger role for PRL in regulating the level of PRLR expression in adipose tissue and liver of male mice compared with female mice.
The size of the retroperitoneal adipose tissue was smaller in the PRL-transgenic females compared with the control mice used in this study. Hyperprolactinemia in humans, especially in males, has been reported to be associated with weight gain (45, 46, 47). However, a decreased weight of adipose tissue was detected in male mice with chronic hyperprolactinemia in parallel with a total body weight increase (11).
In summary, the PRLR has been identified in mouse adipose tissue and adipocytes. The level of PRLR expression increased in adipose tissue during lactation, whereas the level of PRLR mRNA was similar in pregnant and virgin mice. In contrast, PRLR gene expression in liver was higher during pregnancy compared with those in virgin and lactating mice. PRLR gene expression was increased in both adipose tissue and liver of PRL-transgenic mice, indicating that PRL up-regulates its receptor in these tissues. However, the tissue-specific regulation of PRLR expression during pregnancy and lactation indicates that factors other than PRL are of importance for receptor regulation. Furthermore, the dramatic up-regulation of PRLR gene expression in adipose tissue and liver during lactation and pregnancy, respectively, suggests that PRL exerts different metabolic actions during these conditions.
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Acknowledgments |
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Footnotes |
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Received January 21, 2000.
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