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1 Rowett Research Institute, Greenburn Road, Bucksburn, Aberdeen AB21 9SB, UK
2 Department of Hygiene and Human Nutrition, Agricultural University, Wojska Polskiego 31, 60-624 Poznan, Poland
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(Received 25 May 2004;
accepted after revision 8 September 2004;
first published online 9 September 2004)
Corresponding author H. J. McArdle: Rowett Research Institute, Greenburn Road, Bucksburn, Aberdeen AB21 9SB, UK. Email: h.mcardle{at}rri.sari.ac.uk
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Introduction |
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Iron supplementation in appropriate cases therefore is clearly warranted. However, there is still debate whether supplementation should be universally provided. Additionally, neither the optimal time period nor the dose has been agreed. For example, World Health Organization have recommended supplementation of about 60 mg day–1 to all women from the second trimester onward (Stoltzfus & Dreyfuss, 1998; WHO, 2001). In contrast, Casanueva & Viteri (2003) suggest that 60 mg day–1 in normal women is too high, and results in haemoconcentration, decreased birth-weight and increased prematurity. The US Institute of Medicine (Viteri, 1998) recommends a flexible approach, ranging from no supplementation during the first two trimesters to 120 mg day–1, depending on a variety of parameters.
There is very little information about the mechanisms underpinning the differential effects of dose and timing. It is generally agreed that the mother uses both Fe stores and increased absorption to supply the developing fetus. However, how much each can or will contribute is not known. Nor is it clear how the mother will adapt to lower Fe status.
In order to examine these questions, we have developed a rat model of Fe deficiency during pregnancy. The pups born to Fe-deficient dams are smaller than control neonates, and have proportionately smaller livers. However, the degree of Fe deficiency in these pups is less than might be predicted. We have shown that the placenta up-regulates some of the proteins of Fe transport as a response to maternal deficiency. For example, transferrin receptor (TfR) levels increase markedly as deficiency increases (Gambling et al. 2001). Iron is transferred from the transferrin receptor in the endocytotic vesicle into the cell through a channel called divalent metal transporter 1 (DMT1). There are at least two isoforms of this protein. One is regulated by an iron-responsive element (IRE) and the other is not. The IRE form of the protein is also up-regulated in the placenta by maternal Fe deficiency. Consequently, the fetus accumulates Fe at the expense of its mother. How this is reflected in fetal liver metabolism of Fe is part of the subject of this paper. Additionally, we do not yet know whether there is an optimum or preferable time for supplementation. In this paper, therefore, we examine the effect of different time periods of supplementation on growth and development of the offspring.
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The experimental diets were based on a dried egg albumin diet and conformed to American Institute of Nutrition guidelines for laboratory animals (Williams & Mills, 1970). Iron sulphate was added to achieve levels of added Fe of 50 (control diet), 7.5 (Fe-deficient diet) or 75 mg kg–1 (Fe-supplemented diet). Dietary ingredients were purchased from Mayjex Ltd (Chalfont-St Peter, UK), BDH Chemicals (Poole, UK) or Sigma (Poole, UK).
Experimental animals
Experiment was performed using 40 weanling female rats of the Rowett Hooded Lister strain. Animals were housed in cages under constant conditions (temperature 22°C; humidity 55%; 12 h:12 h light:dark illumination photoperiod). All the animals were fed control diet (50 mg Fe (kg diet)–1) for 2 weeks, before being randomized into two groups. The first group, eight animals, remained on the control diet throughout the experiment, including during pregnancy, whilst the remaining 32 animals were placed on the Fe-deficient diet (7.5 mg Fe (kg diet)–1) for 4 weeks before mating. All the rats were mated with males of the same strain. Mating was confirmed by detection of a vaginal plug, and this day was denoted as day 0.5. At 0.5, 7.5 and 14.5 days gestation, separate groups of eight rats were taken from the deficient diet and given the supplemented diet until parturition. The remaining eight rats continued on the Fe-deficient diet. The pups were killed within 12 h of birth by decapitation. The dams were killed by stunning and cervical cord dislocation. All experimental procedures were approved and conducted in accordance with the UK Animals (Scientific Procedures) Act 1986.
Sample collection
The number of neonates (male and female) was counted. Maternal and neonatal blood samples were collected. Maternal livers as well as neonatal livers, hearts, lungs, kidneys and spleens, taken from six neonates (3 male and 3 female), chosen from each mother at random, were rapidly dissected, weighed and frozen in liquid nitrogen before being stored at –70°C.
Haematological measurements
Maternal and neonatal haematocrit were measured by drawing blood into capillary tubes, which were then centrifuged in a high-speed haematocrit centrifuge (Universal 32R, Hettich; Scientific Laboratory Supplies, Coatbridge, UK) and read in a microhaematocrit reader.
Atomic absorption spectrophotometric analyses
For determination of total Fe content in tissues, the heat-dried liver samples were treated with nitric acid (Ultrapure; Merck, Poole, UK). The total Fe content in these samples was determined by graphite furnace atomic spectrophotometry (Aanalyst 600, Perkin Elmer, Beaconsfield, UK). Standards and quality controls were included as appropriate.
Primers for real-time RT-PCR
Complementary DNA PCR primers for the TfR, DMT1 IRE and non-IRE forms, ferritin H and ferritin L were designed using Primer Express software (version 1.5, Applied Biosystems, Warrington, UK) from the DNA sequences GenBank Accession numbers M58040, AF008439, AF029757, NM_012848 and XM_216824, respectively. The primers were as follows: TfR forward primer 1757–1779 bp, reverse primer 1818–1838 bp; DMT1 IRE forward primer 2168–2188 bp, reverse primer 2228–2247 bp; DMT1 non-IRE form forward primer 1650–1673 bp, reverse primer 1700–1722 bp; ferritin H forward primer 585–606 bp, reverse primer 632–656 bp; and ferritin L forward primer 140–156 bp and reverse primer 14–35 bp. The primer sets had a calculated annealing temperature of 58°C. Primers were ordered from MWG Biotech, Munich, Germany.
DNase treatment
Total RNA was prepared by use of TRI reagent (Helena Biosciences, Sunderland, UK) according to the manufacturer's instructions. To eliminate genomic DNA contamination total RNA samples were treated with ribonuclease (RNase)-free Dnase 1 Amplification Grade (Invitrogen Ltd, Paisley, UK), before complementary DNA synthesis. Ten micrograms of total RNA was added to a total volume of 20 µl containing 2 µl 10 x reaction buffer and 1 µl Dnase 1. The total RNA was incubated for 10 min at 25°C, followed by precipitation of the RNA by adding 5 µl 3 M sodium acetate pH 4.8, 55 µl isopropanol and 30 µl diethyl pyrocarbonate (DEPC)-treated water. RNA was precipitated at –20°C for at least 1 h, then centrifuged, washed in 75% ethanol and the pellet dissolved in DEPC-treated water. RNA concentrations were estimated by Agilent analysis (Agilent 2100 Bioanalyser, Agilent Technologies, Stockport, UK).
Reverse transcription
First strand complementary DNA was synthesized by priming with hexamers using the Taqman RT Reagent Kit (Applied Biosystems, Stockport, UK). Reverse transcription was performed in 20 µl reactions using 200 ng of Dnase-treated RNA. Reverse transcription was performed by addition of 12.3 µl RT mix, such that the final concentration was 1 x buffer, 5.5 mM MgCl2, 2 mM dNTP, 2.5 µM hexamer primer, 8 U RNase inhibitor and 25 U MultiscribeTM RT. The mixture was incubated at 25°C for 10 min, incubated at 48°C for 30 min and heated to 95°C for 5 min, and then finally cooled to 4°C. To determine the presence of contaminating cDNA, reverse transcription was omitted in the cDNA synthesis reaction. A specific product was never detected (data not shown).
Real-time quantitative PCR
Real-time PCR amplification and analysis was performed using a 7700 Sequence Detection System (Applied Biosystems) and ABI prism software version 1.9 (Applied Biosystems). Reactions were performed in 25 µl final volume with 300 nM primers and 5 µl cDNA. Magnesium chloride, nucleotides, buffer and Taq DNA polymerase were included in the SYBR Green Master Mix (Applied Biosystems). The PCR amplification was performed according to the manufacturer's instructions and included heating to 50°C for 5 min and denaturation at 95°C for 10 min, followed by 40 cycles with 95°C for 15 s and 60°C for 1 min. To confirm amplification specificity the PCR products from each primer pair were subjected to gel electrophoreses (2.5% Agarose-1000, Invitrogen, with ethidium bromide). For all primer pairs only one product of correct size was detected (data not shown).
Standard curves were generated from increasing amounts of cDNA made from maternal liver control RNA. The cycle threshold (CT) values were used to calculate and plot a linear regression line by plotting the logarithm of template concentration (x axis) against the CT value (y axis). These regression lines were used to calculate the expression level (nanograms of total RNA) for unknown samples.
Statistical analyses
For each dam, the data from the the pups were averaged and recorded as a single point. One-way ANOVA was used to determine statistical significance between multiple data sets and verified by the least significant difference (LSD) test. Linear regression test was used to determine statistical significance between continuous variables. Significance was assumed at P = 0.05. All analyses were carried out using Statistica 5.0 (Statsoft, Polska, Krakow, Poland). The results are presented as means ± S.E.M.
The change in expression in mRNA levels between the control group and the treated groups is presented as a percentage of the control, with the average of the controls set to 100%. Each group contained amplified cDNAs from three plates, and the average of these measurements was used to calculate group mean and S.E.M. Significant differences (P < 0.05) between treated and control groups were determined using Student's unpaired t test, two tailed (GraphPad Instat, version 3.05).
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The results of the present study also confirm our previous findings (Gambling et al. 2002) that neonatal body weight decreases with the severity of maternal Fe deficiency. Fetal growth restriction resulting from severe Fe deficiency has also been demonstrated by Crowe et al. (1995) in rats and Malhotra et al. (2002) in humans.
As might be expected, supplementation reverses the effect. The timing is important, however. Our data show that supplementation in the last third of pregnancy is not as effective as it is when given earlier. Although it is difficult to extrapolate directly from rats to humans, given the differences in timing of developmental milestones, the data support results published in human studies. For example, Hamalainen et al. (2003) demonstrated that maternal anaemia detected in the first trimester was associated with low-birth-weight infants, whereas the mid- and third-trimester anaemia groups did not show significantly different outcomes when compared with non-anaemic women. Additionally, Cogswell et al. (2003) have presented data suggesting that Fe supplementation, even in women of normal Fe status, during the first 28 weeks of pregnancy, but not during the latter half, results in a very significant increase in birth weight.
Even though the rat model has limitations, it also provides us with the possibility of identifying which organs are most susceptible to Fe deficiency. For example, it would seem, from our data, that pre- and early postimplantation embryos are most sensitive to maternal nutritional status. These developmental events occur proportionately much earlier in human pregnancy. Consequently, further studies are needed to identify the developmental windows, rather than the chronological windows, where Fe deficiency exerts its effects.
The distribution of Fe following supplementation is intriguing. The fact that fetal Fe levels recover before that of the mother indicates that Fe is preferentially diverted to the fetus as soon as it becomes available. The mechanism for this can be deduced from this and from our previous studies. Expression of the proteins of Fe transfer is significantly increased in the placenta during Fe deficiency (Gambling et al. 2002, 2003). The degree of increase is significantly greater than that seen in the maternal liver. Clearly, since the present study was carried out on neonates, we have no information on the degree of changes in TfR expression in the placenta. However, assuming they follow the same pattern as previously described, the Fe taken up in the supplemented animals will have been preferentially delivered to the fetus. This is substantiated because there is no significant difference in neonatal expression of the TfR in the liver. In the mother, in contrast, Fe levels did not returned to normal even following 2 weeks of supplementation.
In one sense, the results are not surprising. Iron is critical for rapidly developing fetal and neonatal organ systems. Iron is prioritized to haemoglobin synthesis in red blood cells when Fe supply does not meet Fe demand. Therefore, non-haem-containing tissues such as skeletal muscle, heart and brain will become Fe deficient before signs of Fe deficiency anaemia (Rao & Georgieff, 2002).
The effects on the two forms of DMT1 are also interesting. There are data to suggest that regulation of DMT1 by Fe is altered at different stages in gestation. Lonnerdal and colleagues have shown that there is an apparent developmental regulation of DMT1 in rats (Leong et al. 2003), with greater sensitivity in the gut occurring at postnatal day 20 rather than day 10. In one sense, this is not surprising, since the duodenum is not exposed to the Fe-deficient environment directly until birth. In contrast, as we have previously shown (Gambling et al. 2001) and confirmed in the present study and in Lonnerdal's paper (Leong et al. 2003), liver DMT1 levels are increased in Fe deficiency in the pre- and early postnatal period.
The effect on the non-IRE-regulated DMT1 has not previously been reported. The function of this transcript is not clear, and at present we have no obvious explanation. Ke et al. (2003) have suggested that the two isoforms both respond to Fe deficiency in the heart, while Tchernitchko et al. (2002) suggest that there are other promoters and/or regulators involved in induction of the different isoforms. Clearly, there is much yet to be elucidated.
The effect on ferritin mRNAs was also intriguing. There was no significant change in the maternal liver, or in L ferritin in the neonate, while the H ferritin level significantly decreased both in neonates born from Fe-deficient mothers and in those born from mothers supplemented for 1 week only. H and L ferritin are independently regulated proteins with both transcriptional and translational regulation in response to cellular Fe level. Han et al. (2000) studied the H:L ferritin ratio and mRNA levels in various regions of the brain of male Fe-deficient, control and Fe-supplemented rats. They found that ferritin H and L subunits within the brain respond differently to Fe status. On the basis of experiments with synthetic ferritin heteropolymers, Levi et al. (1994) concluded that the ferritins with high L:H chain ratios are the most efficient in incorporating Fe (Levenson & Fitch, 2000). The data we have obtained in this study would seem to support that observation.
What is most surprising is that there is no correlation between neonatal Fe levels and neonatal size. Instead, the relationship appears to between maternal Fe and the neonate. This suggests that the mother rather than factors deriving from the fetus determines size at birth. There are other data to support these observations. In sheep, Wallace et al. (2002) have shown that placental growth is regulated by maternal nutrition and this, in turn, regulates fetal growth. Various hormone treatments, such as parathyroid hormone-related protein in rats (Wlodek et al. 2004), dexamethasone in sheep (Kerzner et al. 2002) or leptin in sheep (Thomas et al. 2001) have all been shown to alter fetal growth (see Reynolds et al. 2004 for a comprehensive review). At this stage, we are not certain which hormone mediates the effect in anaemic rats, but it is clear that the data may have considerable relevance in understanding the relationship between maternal anaemia and pregnancy outcome in humans.
In summary, therefore, our data show that the developing fetus is very sensitive to maternal Fe status, and that the period of supplementation is critical in reversing the effects of maternal anaemia. Anaemia during pregnancy is endemic in many parts of the world and is a significant concern even in developed countries. It is well established that supplementation is of limited value outside of well-controlled experimental studies. Our data may provide an explanation for this, in that if it is given at an inappropriate time, it will be less effective at reversing the consequences of maternal anaemia. Certainly the data would suggest that early supplementation will be more efficacious than during the last trimester. The mechanisms underlying these observations clearly remain to be determined, but a better understanding will be very valuable in developing the appropriate strategies for treatment of mothers suffering from anaemia during pregnancy.
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References |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Casanueva
E
&
Viteri
FE (2003). Iron and oxidative stress in pregnancy. J Nutr
133, 1700S–1708S.
Cogswell
ME, Parvanta
I, Ickes
L, Yip
R
&
Brittenham
GM (2003). Iron supplementation during pregnancy, anemia, and birth weight: a randomized controlled trial. Am J Clin Nutr
78, 773–781.
Crowe C, Dandeker P, Fox M, Dhingra K, Bennet L & Hanson MA (1995). The effects of anaemia on heart, placenta and body weight and blood pressure in fetal and neonatal rats. J Physiol 488, 515–519.[Medline]
Gambling
L, Charnia
Z, Hannah
L, Antipatis
C, Lea
RA
&
McArdle
HJ (2002). Effect of iron deficiency on placental cytokine expression and fetal growth in the pregnant rat. Biol Reprod
66, 516–523.
Gambling L, Danzeisen R, Gair S, Lea RG, Charania Z, Solanky N et al. (2001). Effect of iron deficiency on placental transfer of iron and expression of iron transport proteins in vivo and in vitro. Biochem J 356, 883–889.[CrossRef][Medline]
Gambling
L, Dunford
S, Wallace
DI, Zuur
G, Solanky
N, Srai
SKS
&
McArdle
HJ (2003). Iron deficiency during pregnancy affects post-natal blood pressure in the rat. J Physiol
552, 603–610.
Hamalainen H, Hakkarainen K & Heinonen S (2003). Anaemia in the first but not in the second or third trimester is a risk factor for low birth weight. Clin Nutr 22, 271–275.[CrossRef][Medline]
Han J, Day JR, Thomson K, Connor JR & Beard JL (2000). Iron deficiency alters H- and L-ferritin expression in rat brain. Cell Mol Biol 46, 517–528.[Medline]
Kapil U & Bhavna A (2002). Adverse effects of poor micronutrient status during childhood and adolescence. Nutr Rev 60, S84–S90.[CrossRef][Medline]
Ke Y, Chen YY, Chang YZ, Duan XL, Ho KP, Jiang DH et al. (2003). Post-transcriptional expression of DMT1 in the heart of rat. J Cell Physiol 196, 124–130.[CrossRef][Medline]
Kerzner LS, Stonestreet BS, Wu KY, Sadowska G & Malee MP (2002). Antenatal dexamethasone: effect on ovine placental 11beta-hydroxysteroid dehydrogenase type 2 expression and fetal growth. Pediatr Res 52, 706–712.[CrossRef][Medline]
Leong WI, Bowlus C & Tallkvist G & Lonnerdal B (2003). DMT1 and FPN1 expression during infancy: developmental regulation of iron absorption. Am J Physiol 285, G1153–G1161.
Levenson CW & Fitch CA (2000). Effect of altered thyroid hormone status on rat brain ferritin H and ferritin L mRNA during postnatal development. Brain Res Dev Brain Res 119, 105–109.[CrossRef][Medline]
Levi S, Santambrogio P, Cozzi A, Rovida E, Corsi B, Tamborini E et al. (1994). The role of the L-chain in ferritin iron incorporation. Studies of homo and heteropolymers. J Mol Biol 238, 649–654.[CrossRef][Medline]
Lewis RM, Forhead AJ, Petry CJ, Ozanne S & Hales CN (2002). Long-term programming of blood pressure by maternal dietary iron restriction in the rat. Br J Nutr 88, 283–290.[CrossRef][Medline]
Lisle SJ, Lewis RM, Petry CJ, Ozanne SE, Hales CN & Forhead AJ (2003). Effect of maternal iron restriction during pregnancy on renal morphology in the adult rat offspring. Br J Nutr 90, 33–39.[CrossRef][Medline]
Malhotra M, Sharma JB, Batra S, Sharma S, Murthy NS & Arora R (2002). Maternal and perinatal outcome in varying degrees of anaemia. Int J Gynaecol Obstet 79, 93–100.[CrossRef][Medline]
Rao R & Georgieff MK (2002). Perinatal aspects of iron metabolism. Acta Paediatr 91 (suppl.), 24–129.
Reynolds LP, Borowicz PP, Vonnahme KA, Johnson ML, Grazuk-Bilska T, Wallace JM et al. (2004). Animal models of placental angiogenesis. Placenta (in press).
Scholl TO & Hediger ML (1994). Anaemia and iron-deficiency anaemia: complication of data on pregnancy outcome. Am J Clin Nutr 59, S492–S500.
Stoltzfus RJ & Dreyfuss ML (1998). Guidelines for the Use of Iron Supplementation to Prevent and Treat Iron Deficiency Anemia. International Nutritional Anaemia Consultative Group (INACG). ILSI Press, Washington, DC, USA.
Tchernitchko D, Bourgeois M, Martin ME & Beaumont C (2002). Expression of the two mRNA isoforms of the iron transporter Nramp2/DMT1 in mice and the funciton of the iron responsive element. Biochem J 363, 449–455.[CrossRef][Medline]
Thomas L, Wallace JM, Aitken RP, Mercer JG, Trayhurn P & Hoggard N (2001). Circulating leptin during ovine pregnancy in relation to maternal nutrition, body composition and pregnancy outcome. J Endocrinol 169, 465–476.[Abstract]
Viteri FE (1998). Prevention of iron deficiency. In Prevention of Micronutrient Deficiencies: Tools for Policymakers and Public Health Workers, ed. Howson CP, Kennedy ET & Horwitz A., pp. 45–102, National Academic Press, Washington, DC, USA.
Viteri FE & Gonzalez MD (2002). Adverse outcomes of poor micronutrient status in children and adolescence. Nutrition Rev 60, S77–S83.[CrossRef][Medline]
Wallace JM, Bourke DA, Aitken RP, Milne JS & Hay WW Jr (2002). Placental glucose transport in growth-restricted pregnancies induced by overnurishing adolescent sheep. J Physiol 547, 85–94.[CrossRef][Medline]
WHO (2001). Iron deficiency anaemia. http://www.who.int/nut/ida.htm.
Williams RB & Mills CF (1970). The experimental production of zinc deficiency in the rat. Br J Nutr 24, 989–1003.[CrossRef][Medline]
Wlodek ME, Di Nicolantonio R, Westcott KT, Farrugia W, Ho PW & Moseley JM (2004). PTH/PTHrP receptor and mid-molecule PTHrP regulation of intrauterine PTHrP: PTH/PTHrP receptor antagonism increases SHR fetal weight. Placenta 25, 53–61.[CrossRef][Medline]
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) and IRE-regulated forms (
) were measured. Samples were taken and mRNA prepared, reverse transcribed and measured, as described in the Methods. The results are the means ±
S.E.M. of livers from at least 6 dams and 6 offspring, each taken from a different litter. *P < 0.05; **P < 0.01 when compared to the control value by Student's unpaired t test.
