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Endocrinology Vol. 142, No. 2 532-537
Copyright © 2001 by The Endocrine Society

ARTICLES

Body Weight and Fat Deposition in Prolactin Receptor-Deficient Mice1

Michael Freemark, Don Fleenor, Phyllis Driscoll, Nadine Binart and Paul A. Kelly

Departments of Pediatrics (M.F., D.F., P.D.) and Cell Biology (M.F.), Duke University Medical Center, Durham, North Carolina 27710; and INSERM U-344, Faculté de Médecine Necker, Paris, France

Address all correspondence and requests for reprints to: Dr. Michael Freemark, Department of Pediatrics, Box 3080, Duke University Medical Center, Durham, North Carolina 27710. E-mail: freem001{at}mc.duke.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
To explore the roles of the lactogens in adipose tissue development and function, we measured body weight, abdominal fat content, and plasma leptin concentrations in a unique model of lactogen resistance: the PRL receptor (PRLR)-deficient mouse. The absence of PRLRs in knockout mice was accompanied by a small (5–12%), but progressive, reduction in body weight after 16 weeks of age. Females were affected to a greater degree than males. The reduction in weight in female PRLR-deficient mice (age 8–9 months) was associated with a 49% reduction in total abdominal fat mass and a 29% reduction in fat mass expressed as a percentage of body weight. Lesser reductions were noted in male mice. Plasma leptin concentrations were reduced in females but not in males. That the reductions in abdominal fat may reflect in part the absence of lactogen action in the adipocyte is suggested by the demonstration of PRLR messenger RNA in normal mouse white adipose tissue. Nevertheless, steady state levels of PRLR messenger RNA in mature adipocytes are very low, suggesting that the effects of lactogens might be mediated by other hormones or cellular growth factors. Our observations suggest roles for the lactogens in adipose tissue growth and metabolism in pregnancy and postnatal life.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE LACTOGENIC hormones PRL and placental lactogen (PL) play roles in carbohydrate metabolism through effects on pancreatic insulin production and peripheral insulin sensitivity (1, 2, 3, 4, 5, 6, 7, 8). However, the roles of the lactogens in lipid metabolism are poorly understood. Two lines of evidence suggest that the lactogens exert lipolytic actions in white adipose tissue during lactation and late pregnancy. First, the hyperprolactinemia of lactation is accompanied by depletion of abdominal fat stores and synthesis and secretion of triglycerides by the mammary gland (9, 10), and second, hPL stimulates lipolysis in adipose tissue in vivo and in vitro and potentiates the lipolytic effects of theophyline (11, 12, 13, 14, 15, 16, 17, 18, 19). These observations must be interpreted with caution, however, because 1) a reduction of circulating PRL concentrations in lactating rats has no effect on abdominal lipid synthesis and storage (10), and 2) the hPL used for early investigations was purified from human placenta and may have been contaminated by trace amounts of placental GH, a potent lipolytic agent. In addition, under some experimental conditions hPL stimulates glucose uptake and glycogen synthesis in isolated adipocytes, mimicking the lipogenic effects of insulin (20).

More importantly, recent investigations have failed to demonstrate lipolytic effects of ovine PL or recombinant bovine PL, bovine PRL, mouse PRL, or mouse PL in homologous systems (21, 22, 23). Indeed, PRL stimulates food intake and fat deposition in female rats (24, 25, 26) and birds (27, 28, 29) and has lipogenic effects in fetal and newborn rat hepatocytes (30, 31).

To clarify the roles of the lactogens in fuel homeostasis, we measured body weight, abdominal fat content, and plasma leptin in a unique model of lactogen resistance: the PRL receptor (PRLR)-deficient mouse. This experimental model was created by targeted deletion of the gene encoding the mouse PRLR (32). PRLR knockout mice are resistant to the actions of mouse PRL and mouse PL, which bind only to the mouse PRLR. Female homozygous PRLR-deficient mice are sterile, a consequence of progesterone deficiency, hypoestrogenemia, and defects in egg transport and implantation. The male homozygous mutants, on the other hand, appear to have near-normal reproductive capacity and normal serum testosterone concentrations (33). PRLR-deficient mice also have reduced rates of bone formation, decreased bone mineralization, and hyperparathyroidism (33), but the effects of PRLR deficiency on weight gain and fat deposition have not been examined previously.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Mice
The generation of PRLR-deficient mice has been described in detail previously (32, 33). Heterozygous mutants (129Sv/C57BL/6) were bred to produce -/-, +/-, and +/+ animals. The pups were genotyped by PCR amplification of the NEO gene using specific primers, described previously (33). The mode of handling and treatment of laboratory mice were approved by the institutional committee on the treatment of laboratory animals of Duke University Medical Center.

The mice were maintained on a 12-h light, 12-h dark cycle (lights on, 0700–1900 h) with food and water provided ad libitum. The chow (Laboratory Rodent Diet 5001, Ralston Purina Co., St. Louis, MO) provided 12.1% of calories as fat, 28% as protein, and 59.8% as carbohydrate. All mice were virginal and were housed in cages in groups of four or five. Body weights of individual mice were measured between 12–75 weeks of age using a cross-sectional approach. A single PRLR-deficient male that weighed 26 g at 68 weeks of age was eliminated as an outlier before final analysis of growth data.

Blood samples were secured at 0900 h after a 17-h fast. The blood was obtained rapidly (15–20 sec) by retroorbital puncture without anesthesia and was collected into EDTA-coated tubes. All animals were sampled on at least three separate occasions between 6 and 9 months of age; plasma leptin levels were analyzed on each of the samples in separate assays. Plasma leptin was measured using mouse RIA kits purchased from Linco Research, Inc. (St. Louis MO). The mice were killed by cervical dislocation. Abdominal fat stores were measured at the time of death; all adipose tissue that was grossly visible was retrieved and weighed.

RT-PCR and Southern analysis
Samples of adipose tissue were obtained from wild-type males (M), virgin female (F), and lactating females (LF). Three animals were analyzed from each group, and the experiments were repeated three times.

Total RNA was prepared from abdominal fat of adult mice using Tri-Reagent (Molecular Research Center, Inc., Cincinnati, OH) according to the instructions of the manufacturer. The RNA was reverse transcribed into complementary DNA (cDNA) using the following protocol. Five micrograms of total RNA were incubated for 60 min at 37 C with 200 U Moloney murine leukemia virus reverse transcriptase (Life Technologies, Inc./BRL, Gaithersburg, MD) in buffer (20 mM Tris-HCl, pH 8.3, with 50 mM KCl and 5 mM MgCl2) containing 1 mM deoxynucleotide triphosphates (Promega Corp.), 10 mM dithiothreitol, 30 U ribonuclease inhibitor (RNAsin), and 1 µg random hexamer oligonucleotides in a total volume of 30 µl. Control samples contained either no RNA or no reverse transcriptase.

One fifth of the cDNA generated under these conditions was subjected to PCR using primers encoding nucleotides 600–619 (5'-GACTCGCTGCAAGCCAGACC-3', sense) and 1018–1037 (5'-TGACCAGAGTCACTGTCAGG-3', antisense) of the long isoform of the mouse PRLR (34). Additional aliquots of cDNA were subjected to PCR using a sense primer (5'-GAGAAAAACACCTATGAATGTC-3', exon 5) common to all forms of the receptor and an antisense primer common to all forms (5'-CGTCTACTCATAGTTTAGGA-3', exon 9) or antisense primers encoding the short isoform PRL-Rs1 (5'-CCTTGAGACTAGATTATTGGC-3', exon 11) or the long isoform (5'-CAATCTGTCCATAAGTCTAGC-3', exon 10). The primer pairs designated 5F-9R, 5F-10R, and 5F-11R refer to the forward primer in exon 5 and the reverse primers in exons 9, 10, and 11. The reaction buffer contained 18.6 mM Tris-HCl, pH 8.3, with 45.9 mM KCl, 3 mM MgCl2, 0.2 mM deoxy-NTPs, 0.2 U Taq polymerase (Life Technologies, Inc./BRL), and 25 pmol of each of the primers in a final volume of 50 µl. After a 4-min denaturation at 94 C, the samples were subjected to 30 cycles of PCR. Samples were denatured at 94 C for 45 sec, annealed at 56-60 C for 45 sec, and extended at 72 C for 1–2 min, with a final elongation cycle at 72 C for 10 min. The samples were separated on a 1.2% agarose gel and transferred to Boeh- ringer-Mannheim-charged membranes. The membranes were then probed with a digoxigenin-labeled RNA probe encoding bases 553-1191 of the mouse PRLR. The probe was generated using a BamHI-EcoRI fragment of a plasmid containing the mPRLR (pSP73-PRL-Rs1, provided by Dr. Daniel Linzer). The antisense RNA was generated using T3 RNA polymerase and was labeled with digoxigenin using a kit from Roche Molecular Biochemicals (Indianapolis, IN).

The membranes were incubated at 42 C overnight in a hybridization mixture containing 5 x SSC (standard saline citrate), 0.1% N-lauroylsarcosine, 0.02% SDS, 1% blocking reagent (Roche Molecular Biochemicals), and 50% formamide. On the following morning the membranes were washed twice in 2 x SSC with 0.1% SDS (15 min each), twice with 0.5 x SSC with 0.1% SDS, and twice with 0.1 x SSC with 0.1% SDS (10 min). All washes were performed at 68 C. After rinsing with malate buffer, the membranes were incubated in blocking solution for 1 h and then with antidigoxigenin-alkaline phosphatase (1:5000) for 30 min. The membranes were washed in malate buffer three times for 15 min each time. Chemiluminescent detection of the digoxigenin-labeled probe was performed by applying CSPD (10 µg/ml; Tropix, Bedford, MA) diluted in buffer 3 [100 mM Tris-HCl (pH 9.5), 100 mM NaCl, and 1 mM MgCl2] and exposing the membranes to Hyperfilm enhanced chemiluminescence (Amersham Pharmacia Biotech, Elk Grove, IL) for 5 min.

Statistical analysis
Differences among sample means were assessed by ANOVA followed by the Newman-Keuls test of multiple comparisons. All experiments were repeated on at least two or three occasions. P < 0.05 was considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
In a cross-sectional analysis, the weights of PRLR-deficient mice were compared with the weights of wild-type littermate controls between 12 and 76 weeks of age. As shown in Fig. 1Go, the weights of the PRLR-deficient males and females were comparable to those of their wild-type littermates at 12–16 weeks of age. After 16 weeks of age, however, the weight curves of the PRLR-deficient mice and their wild-type controls appeared to diverge. Between 16 and 76 weeks, the slopes of the regression lines defining weight gain in the PRLR-deficient males (0.12) and females (0.06) were significantly less than the slopes of the regression lines depicting weight gain in the wild-type animals (0.20–0.21). This finding suggests that the rates of weight gain in wild-type mice after 16 weeks of age exceed the rates of weight gain in PRLR-deficient mice.



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  		Figure 1. Cross-sectional analysis of body weights of PRLR-deficient (-/-) and wild-type (+/+) mice. Individual male (+/+, n = 37; -/- n = 35) and female (+/+, n = 78; -/-, n = 47) mice were weighed in the morning at the ages indicated on the graph. Equations represent the lines generated by regression analysis, with y representing weight in grams and x representing weeks of age.

 
The differences in weight between PRLR-deficient and wild-type mice were more conspicuous in females than in males. At 27–37 weeks of age, the weights of homozygous mutant males (34.3 ± 0.8 g; n = 12) were not significantly different from the weights of wild-type males (35.8 ± 1.3 g; n = 7). At that stage, however, homozygous mutant females (25.2 ± 0.5 g; n = 19) weighed 8.3% less (P < 0.01) less than wild-type females (27.8 ± 0.5 g; n = 31). By 42–76 weeks of age the PRLR-deficient males (36.1 ± 0.8 g; n = 13) weighed approximately 12% less (P < 0.03) than wild-type males (41.1 ± 1.0 g; n = 19), whereas PRLR-deficient females (27.4 ± 0.9 g; n = 13) weighed 12.3% less (P < 0.01) than wild-type females (31.1 ± 0.9 g; n = 12). It should be noted that there was considerable variability in the weights of wild-type and PRLR-deficient mice at all developmental stages.

The decrement in body weight in adult PRLR-deficient mice (age, 8–9 months) was accompanied by a reduction in abdominal fat stores. Total abdominal fat content, reflecting sc, mesenteric, perirenal, and epididymal (male) or periovarian (female) fat stores, was 34% less (P < 0.05; Fig. 2Go left) in mutant males (1.05 ± 0.20 g; n = 11) than in wild-type males (1.58 ± 0.18 g; n = 12) and 49% less (P < 0.02; Fig. 2Go, left) in mutant females (0.67 ± 0.14 g; n = 9) than in wild-type females (1.31 ± 0.22 g; n = 7). Significant differences between PRLR-deficient and wild-type females were also detected when total abdominal fat was expressed as a function of total body weight (+/+ females, 4.5 ± 0.4%; -/- females, 3.2 ± 0.3%; Fig. 2Go, right).



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  		Figure 2. Total abdominal fat content of PRLR-deficient (-/-) and wild-type (+/+) mice at 8–9 months of age. Abdominal fat content represents stores of sc, mesenteric, perirenal, and epididymal (male) or periovarian (female) fat. +/+ males, n = 12; -/- males, n = 11; +/+ females, n = 10; -/- females, n = 11. Left, Absolute values; right, expressed as a percentage of body weight.

 
The reductions in abdominal fat content in PRLR-deficient females were accompanied by a 40% reduction in fasting plasma levels of the adipocyte hormone leptin (P < 0.02; Fig. 3Go). Leptin levels in PRLR-deficient males did not differ from those in wild-type littermates. A reduction in plasma leptin in female mice was also noted when leptin levels were expressed as a function of body weight [+/+ females, 0.26 ± 0.04 ng/ml·g; -/- females, 0.17 ± 0.02 ng/ml·g (P < 0.05); +/+ males, 0.13 ± 0.03; -/- males, 0.13 ± 0.02].



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  		Figure 3. Plasma leptin concentrations in PRLR-deficient (-/-) and wild-type (+/+) mice at 7–9 months of age. Blood samples were obtained after a 17-h overnight fast. +/+ males, n = 11; -/- males, n = 9; +/+ females, n = 12; -/- females, n = 14. Similar results were noted in three separate experiments.

 
To determine whether the effects of PRLR deficiency on adipose tissue mass might be mediated directly by changes in PRL action in the adipocyte, we examined the expression of PRLR messenger RNA (mRNA) in mouse adipose tissue. As shown in Fig. 4Go, A and B, the mRNAs encoding the long and short isoforms of the PRLR are expressed in adipose tissue of normal male mice, virgin female mice, and lactating female mice; PCR of adipose tissue cDNA generated a single product that hybridized to a digoxigenin-labeled RNA probe encoding the mouse PRLR. However, the levels of adipose tissue PRLR mRNA, as determined by Northern analysis, are very low, near the limits of detectability (not shown). This observation concords with recent studies demonstrating low levels of PRLR immunoreactivity in mouse adipose tissue (35).



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  		Figure 4. A, mRNA encoding the long isoform of the PRLR is expressed in mouse white adipose tissue. Total RNA from abdominal fat from wild-type adult males (M), adult virgin females (F), and lactating females (LF) was used for the preparation of cDNA. The cDNA was then subjected to RT-PCR using primers encoding the long isoform of the mouse PRLR. Control samples contained no reverse transcriptase (-RT) or no RNA. The single 438-bp product generated by PCR hybridized to a digoxigenin-labeled probe encoding the mouse PRLR. B, mRNA encoding the short as well as the long isoforms of the PRLR are expressed in mouse white adipose tissue. Total RNA from abdominal fat (adult virgin female mice) was used for the preparation of cDNA. The cDNA was then subjected to RT-PCR using primers encoding a product common to both long and short isoforms (5F/9R), to the long isoform (5F/10R), or to the short (5F/11R) isoform of the mouse PRLR. The lane labeled -RT represents the PCR reaction after a control cDNA incubation performed in the absence of reverse transcriptase. The various PCR products are of the expected sizes (common 5F/9R, 569 bp; 5F/10R, 653 bp; 5F/11R, 690 bp) and hybridized to digoxigenin-labeled probes encoding the mouse PRLR (not shown).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The absence of PRLRs in knockout mice was accompanied by a small, but progressive, reduction in the rate of weight gain after 16 weeks of age and a reduction in abdominal fat mass. Females were affected to a greater extent than males. As serum leptin concentrations in humans and rodents correlate strongly with adipocyte mass and percent body fat, the reduction in fat mass in PRLR-deficient females probably contributed to the reduction in plasma leptin levels. Leptin levels in PRLR-deficient males, on the other hand, were not reduced significantly. This may reflect the fact that the relative reductions in fat mass (as a percentage of body weight) in PRLR-deficient males were not as great as the reductions in fat mass in PRLR-deficient females. In addition, leptin levels correlate more strongly with the amount of sc fat than with visceral fat stores (36) and are higher in females than in males. It is possible that the effects of PRLR deficiency on the distribution of fat in females may differ from its effects on fat distribution in males.

Our findings of reduced fat content and leptin deficiency in PRLR-deficient mice would seem to contradict prevailing hypotheses that ascribe to the lactogens a major role in the accentuated lipolysis of late pregnancy. Nevertheless, the literature provides evidence that lactogenic hormones may have lipogenic as well as lipolytic effects in vivo. First, PRL stimulates fat deposition and weight gain in female rats, pigeons, ring doves, and sparrows and may contribute to the seasonal fattening of birds in preparation for flight (24, 27, 28, 29). Second, PRL stimulates increases in white adipose tissue leptin mRNA and plasma leptin levels in female rats in vivo (37). Finally, available (albeit limited) evidence suggests that hyperprolactinemia in men and nonpregnant women may be accompanied by weight gain (38, 39, 40, 41). The weight gain in men may be reversed by bromocryptine, which reduces serum PRL levels (40). Bromocryptine also reduces food intake, body fat, hepatic triglyceride synthesis, and adipose tissue lipoprotein lipase activity in obese mice (42, 43). Notwithstanding the fact that bromocryptine may have independent effects on carbohydrate and lipid metabolism, these observations suggest that lactogens may have adipogenic activity in vivo, at least under certain conditions.

The association of hyperprolactinemia with abdominal fat deposition does not pertain during lactation, when abdominal fat stores are depleted, and triglycerides are synthesized and secreted by the mammary gland (18, 19). The roles of PRL and other hormones in the conversion from lipogenesis to lipolysis in abdominal fat are unclear. Insulin action in adipose tissue is blunted during lactation (44), but this is not induced by hyperprolactinemia because 1) bromocryptine treatment of lactating rats reduces serum PRL concentrations, but has no effect on lipid synthesis in abdominal fat; and 2) a reduction in serum PRL attenuates the lipogenic response to insulin (19, 44). Williamson and his colleagues conclude that the reduction in adipose tissue stores during lactation reflects a resistance to insulin action rather than the increase in serum PRL levels. It is possible that the effects of lactogens on adipose tissue metabolism may vary with developmental stage and with changes in the prevailing hormonal environment. It should be noted that our measurements of abdominal fat content were performed only in adult mice; it is possible that the effects of PRLR deficiency on abdominal fat might vary with age and pubertal status.

What explains the changes in body weight, abdominal fat content, and serum leptin in adult PRLR-deficient mice? There are at least four possible contributing factors. First, as lactogenic hormones stimulate food intake in pigeons, ring doves, sparrows, sheep, and female rats (24, 25, 26, 27, 28, 29, 45), the reduction in body weight and fat content of PRLR-deficient mice may reflect in part a reduction in caloric intake. Second, the reduction in abdominal fat content of PRLR-deficient mice may reflect a reduction in insulin production, as suggested by studies of the effects of lactogenic hormones in isolated pancreatic islets and in rat insulinoma cells (4, 5, 6, 7, 8). Third, PRLR deficiency in female mice is accompanied by a state of progesterone deficiency and hypoestrogenemia (32, 33). As progesterone stimulates food intake and fat deposition (46, 47), and estrogen induces leptin production in female rats (48, 49), the reduction in abdominal fat content and serum leptin in female PRLR-deficient mice may derive in part from a deficiency of sex steroids. Interestingly, the testosterone levels in male PRLR-deficient mice are normal (33); thus, the small reductions in abdominal fat content in male mice cannot be explained by a deficiency or an excess of testosterone.

Finally, the detection of PRLRs in normal mouse adipose tissue (this study and Ref. 35) and in the brown and white adipose tissue of fetal rats and sheep (50, 51, 52) suggests that the reductions in abdominal fat in PRLR-deficient mice might reflect the failure of lactogens to exert direct lipogenic effects on adipose cell development and/or metabolism. PRLR expression is induced during the differentiation of adipocytes from primary mouse bone marrow stromal cells (53), and PRL stimulates adipogenic conversion of NIH-3T3 preadipocytes and enhances their expression of adipocyte genes, including CCAAT enhancer-binding protein-ß, peroxisome proliferator-activated receptor-{gamma}, adipsin, and lipoprotein lipase (54, 55). Although the levels of PRLR mRNA and PRLR immunoreactivity (35) in mature mouse adipocytes are quite low, the expression of PRLRs in preadipocytes has not been examined systematically. Thus, it remains unclear whether the effects of lactogens on adipose tissue development or function are mediated directly through effects on the adipocyte or preadipocyte or indirectly through effects on hypothalamic function or through changes in the production, secretion, or action of other hormones or growth factors, such as the glucocorticoids, thyroid hormone, the catecholamines, GH, and the insulin-like growth factors.

Although we found a small reduction in weight gain in PRLR-deficient mice, no apparent deficit in weight gain was reported in mice with a targeted deletion of the gene for PRL (56, 57). The PRL-deficient mice were assessed at 2–6 weeks and at 6 months of age. Given the progressive reduction in weight gain with age in the PRLR-deficient mice, it is possible that deficits in weight gain in PRL-deficient mice might not emerge until after 6 months of age. Alternatively, the phenotypes of the PRLR-deficient and PRL-deficient mice may differ in certain respects. Phenotypic differences between the PRLR-deficient and the PRL-deficient mice might be related in part to differences in exposure to lactogenic hormones in utero. Mouse PRL is not detected in serum until after birth (58), but mouse PL II circulates in fetal serum in mid- to late gestation (59). PRLR-deficient mice, being resistant to the effects of PL as well as PRL, are deprived of the biological actions of mouse PL II during fetal life. In contrast, PRL-deficient mice are theoretically exposed to PL II in utero, although the levels of PL II in fetal PRL-deficient mice have not yet been measured.

Novel roles for the lactogens in fetal and maternal adipose tissue development and function are suggested by the patterns of hormone production, adipose tissue accumulation, and leptin expression during pregnancy. The mass of adipose tissue and the serum concentrations of leptin increase during the first 26–32 weeks of gestation in the pregnant mother and during the third trimester in the human fetus (60, 61, 62, 63, 64). The accumulation of fat mass and the rise in serum leptin coincide with striking increases in the concentrations of lactogenic hormones in maternal and fetal blood. In pregnant women, for example, the concentrations of hPL and PRL increase progressively between 10 and 36 weeks gestation to levels approximating 6000 and 130 ng/ml, respectively (65). In the human fetus, serum hPL concentrations rise from 5 ng/ml at midgestation to approximately 30 ng/ml at term (66), whereas PRL concentrations increase exponentially from 10–20 ng/ml at 28 weeks to 150–300 ng/ml at term (67). Thus, increases in the lactogenic hormones together with high levels of progesterone and estrogen (46, 47, 48, 49, 68) may contribute to the accumulation of adipose tissue stores and induction of serum leptin in the mother and fetus. Interestingly, the percent body fat mass and the concentrations of leptin in pregnant women decline slightly during the third trimester (62, 63, 64). This may reflect the rising levels of placental GH, which has potent lipolytic effects and reduces serum leptin concentrations in vivo (69, 70, 71). Fat mass and leptin concentrations may not decline in the human fetus in late gestation because placental GH does not circulate in fetal serum (69) and because there is a relative deficiency of GH receptors in fetal tissues (65).

In summary, the loss of PRLRs is associated with reductions in body weight and abdominal fat mass and hypoleptinemia in females. These observations suggest novel roles for the lactogens in adipose tissue development and function during pregnancy and postnatal life.


   Acknowledgments
 
The authors thank Dr. Chris Ormandy for helpful comments.


   Footnotes
 
1 This work was supported in part by grants from the NICHD (HD-24192 to M.F.), the Juvenile Diabetes Foundation (196029 to M.F.), Eli Lilly & Co. (to M.F.), and INSERM (to P.A.K.). Back

Received August 18, 2000.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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