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The World Health Organization considers iron deficiency the number one nutritional disorder in the world.

This is a liquid 1 oz dropper bottle of a Mastery for your Blood Magnetism!!!!!

This is another Pre-Order Sale!!!! Product Ships 10-26-17 and yes its already in stock so don’t wait!!!!


When we have a good bloodstream we have a good, healthy life. It would be wise to take time to learn the principles of building good blood in the body. Carbon dioxide and other waste gases are re-absorbed into the life-giving oxygen. Everyone knows that two atoms of oxygen unite with one atom of carbon to form dioxide. But when there is insufficient oxygen, only one atom unites with carbon, to produce carbon-monoxide, and that is where most of our trouble begins–anemia, low blood pressure, or where there is an abundance of calcium, high blood pressure; because calcium thickens the blood

It requires a great deal more pressure to pump thick blood than it does to pump thin blood; and please make special note, that all this is brought about because there is not sufficient iron in the blood, to carry enough oxygen to the cells, to enable them to breathe, and throw off their waste products. New cells are not produced fast enough to replace the decaying and dead ones. 

Pus is formed only when cells decay. Therefore, it requires no great stretch of the imagination to see how vitally necessary it is to have enough iron in the blood stream to convey sufficient oxygen to all parts.

It is impossible for you to get sick if your iron level is up to par. If you have a disease, no matter what kind of disease it is, you are anemic. Iron is the mineral that conveys oxygen to the brain. Iron should have carbon, hydrogen and oxygen. (the CHO chain). Iron is the spark plug of the human body. When you are deficient in iron, you are susceptible to a whole bunch of diseases. Iron fires the body up and is the only mineral on the planet that is magnetic. Being that iron is magnetic it has a tendency to pull other minerals to it.

It pulls magnesium, zinc, gold, calcium, phosphorous, etc. It would be safe to say that when you take large doses of iron that you are taking all the other minerals. The lack of iron causes 40 manifestations of disease. Without iron, the body looses energy and the immune system begins to give way. There is no oxygen going to the brain when iron is low. Without iron, we wrinkle at a young age and no longer able to walk straight. The body naturally consumes or uses the amount of 3 table spoons of iron a day.

It is the deprivation of iron fluorine. Sickle Cell Anemia is when the blood plasma has broken down by mucous into a sickle. Mucous sinks into the plasma, into the cell itself, breaks and disunites the cell. Removing the mucous the cell unites again. To maintain that level, you have to feed the patient large doses of iron phosphate. Not ferrous oxide.

You need iron.

– Dr. Sebi


Iron deficiency in children with mitochondrial disease

Mitochondrial disease is an energy metabolic disorder with various organ involvement. Iron is widely known to be one of the most important nutriments required for normal brain development and several essential metabolic functions. We retrospectively studied the laboratory data on iron deficiency (ID) in 69 children with mitochondrial respiratory chain complex (MRC) defects by biochemical enzyme assay using muscle tissue. We analyzed the differences between groups of mitochondrial disease based on the presence of ID. ID has higher prevalence in children with mitochondrial disease than in the normal population. There were 6 (9%) patients with low hemoglobin, 12 (17%) with low serum ferrtin, and 22 (32%) with low transferrin saturation levels among children with MRC defects. In comparisons between the ID and the non-ID group of MRC-defect patients, the frequency of MRC I defect was significantly higher in the ID group while that of MRC IV defect was higher in the non-ID group. Abnormal brain magnetic resonance imaging (MRI) findings were more frequently detected in the ID group. The incidence of failure to thrive and gastrointestinal symptoms were significantly higher in the ID group. Early diagnosis and proper treatment of ID are recommended. Especially in cases with risk factors such as failure to thrive or gastrointestinal manifestation, active evaluation of ID should be encouraged.

Department of Pediatrics, Gangnam Severance Hospital, Severance Children’s HospitalYonsei University College of MedicineGangnam-guKorea

The Role of Mitochondrial Metabolism and Redox Signaling in Iron Deficiency Anemia

Grant C BullockChanté L RichardsonValerie SchrottNaomi D GunawardenaTeague Nolan ColeCatherine G Corey and Sruti S Shiva


Several clinical observations illustrate the link between iron and erythropoietin (EPO)-mediated signaling in committed erythroid progenitor cells. In iron deficiency anemia (IDA), erythropoiesis is blocked despite increased serum EPO concentrations. Intravenous iron improves the effectiveness of exogenous EPO in patients with EPO-refractory anemia of chronic disease. These clinical observations suggest that iron dominantly regulates EPO-receptor signaling. However, the mechanism of this iron-mediated signaling remains unclear. We recently demonstrated that 1) the aconitases, multifunctional iron-sulfur cluster proteins that convert citrate into isocitrate are essential in the iron- EPO-signaling pathway in erythroid progenitor cells, and that 2) isocitrate, the product of aconitase, can enhance the effectiveness of EPO during iron deficiency in vitro and in mice with IDA and in rats with the anemia of chronic inflammation. These observations suggest that isocitrate, or its derivatives that synergize with erythropoiesis stimulating agents, have important therapeutic application in the treatment of anemia. New data shows that cellular iron restriction regulates mitochondrial oxygen consumption rates (OCR) differentially over time during erythropoiesis, suggesting a novel link between mitochondrial function and erythropoeisis. It is unknown how iron deficiency induced inhibition of mitochondrial aconitase (ACO2) regulates mitochondrial metabolism during RBC production. Pilot data show that ACO2 inhibition by cellular iron deprivation or pharmacological inhibition of ACO2 decreases mitochondrial respiratory rates (RRs) and alters reactive oxygen species (ROS) production. Further, isocitrate normalizes mitochondrial RRs and ROS and restores RBC production. Importantly, disruption of mitochondrial ROS generation with a mitochondrial-specific anti-oxidant blocks RBC production and a subset of oxidant generators promote erythropoiesis. Targeted reduction of ACO2 protein expression and enzyme activity in iron replete stably transduced K562 cells decreases OCRs. This confirms the link between ACO2 and mitochondrial metabolism in human erythroid cells. These data inform our overarching hypothesis that iron-restriction inhibits ACO2, thereby inhibiting mitochondrial metabolism, resulting in the loss of a mitochondrial ROS signal that is required for erythropoiesis. The loss of this critical mitochondrial ROS signal inhibits the EPO signaling that is required for RBC production. These data also suggest that ACO2 is an iron-sensing regulator of mitochondrial metabolism and ROS signaling.

Dietary iron deficiency induces ventricular dilation, mitochondrial ultrastructural aberrations and cytochrome c release: involvement of nitric oxide synthase and protein tyrosine nitration

Feng DongXiaochun ZhangBruce CulverHerbert G. Chew JrRobert O. KelleyJun Ren


Iron deficiency is associated with multiple health problems, including the cardiovascular system. However, the mechanism of action of iron-deficiency-induced cardiovascular damage is unclear. The aim of the present study was to examine the effect of dietary iron deficiency on cardiac ultrastructure, mitochondrial cytochrome c release, NOS (nitric oxide synthase) and several stress-related protein molecules, including protein nitrotyrosine, the p47phox subunit of NADPH oxidase, caveolin-1 and RhoA. Male weanling rats were fed with either control or iron-deficient diets for 12 weeks. Cardiac ultrastructure was examined by transmission electron microscopy. Western blot analysis was used to evaluate cytochrome c, endothelial and inducible NOS, NADPH oxidase, caveolin-1 and RhoA. Protein nitrotyrosine formation was measured by ELISA. Rats fed an iron-deficient diet exhibited increased heart weight and size compared with the control group. Heart width, length and ventricular free wall thickness were similar between the two groups. However, the left ventricular dimension and chamber volume were significantly enhanced in the iron-deficient group compared with controls. Ultrastructural examination revealed mitochondrial swelling and abnormal sarcomere structure in iron-deficient ventricular tissues. Cytochrome c release was significantly enhanced in iron-deficient rats. Protein expression of eNOS (endothelial NOS) and iNOS (inducible NOS), and protein nitrotyrosine formation were significantly elevated in cardiac tissue or mitochondrial extraction from the iron-deficient group. Significantly up-regulated NADPH oxidase, caveolin-1 and RhoA expression were also detected in ventricular tissue of the iron-deficient group. Taken together, these results suggest that dietary iron deficiency may have induced cardiac hypertrophy characterized by aberrant mitochondrial and irregular sarcomere organization, which was accompanied by increased reactive nitrogen species and RhoA expression.


Deficiency in dietary iron intake has been demonstrated to be one of the most prevalent nutritional problems in the world with approximately 5 billion people afflicted. Nutritional iron deficiency is believed to trigger multiple cardiovascular diseases, including cardiac hypertrophy and chronic heart failure [13]. Although the precise mechanism of action responsible for dietary iron-deficiency-induced cardiac dysfunction and morphological aberration remains poorly understood, several adaptive or compensatory mechanisms have been postulated for altered cardiac function and morphology as a result of dietary iron deficiency. For example, erythropoietin production was elevated under iron deficiency and associated with increased intra-erythrocytic levels of 2,3-diphosphoglycerate [4]. In addition, structural remodelling and compensatory vascular dilatation may contribute to decreased systemic peripheral resistance and reduced afterload, which in turn increases stroke volume under iron deficiency [5,6]. Blood viscosity and sympathetic tone were reduced and elevated respectively, under iron-deficiency-induced anaemia, leading to enhanced venous return, preload and heart rate [7]. These haemodynamic and non-haemodynamic changes (e.g. preload, afterload, heart rate, stroke volume and erythropoietin) have validated the significance of iron content in the regulation of cardiovascular function and morphology.

Mitochondria are crucial players for various physiological processes, including embryonic development, fat metabolism and aging. More importantly, mitochondria are essential organelles for the pro- and anti-oxidant balance responsible for the pathogenesis of a number of cardiovascular disorders [8]. Mitochondria dominate oxygen consumption and energy expenditure [9]. On the other hand, iron is a major cofactor for a number of respiratory chain enzymes [10], many of which reside in mitochondria [10]. It is thus natural to speculate that mitochondria may be a target for anaemia/hypoxia induced by dietary iron deficiency. As a potent vasodilator, NO (nitric oxide) regulates tissue blood flow, oxygen supply and respiratory substrates to mitochondria. Recent studies indicated that NO generated by eNOS [endothelial NOS (NO synthase)] and iNOS (inducible NOS) may be the ‘missing link’ in mitochondrial oxidative damage via reaction with O2(superoxide anion) and ONOO (peroxynitrite) formation [9,11]. It is also suggested that the biogenesis of mitochondria is tightly regulated by the levels of eNOS, NO or its second messenger cGMP [9,11]. It could be suggested that the diverse interactions between NO and mitochondria may be linked to oxidative stress and cardiac damage. eNOS and iNOS are key enzymes determining the levels of NO en route to RNS (reactive nitrogen species) injury if overproduced [12]. Activation of NADPH oxidase generates O2, which rapidly reacts with NO to form ONOO, one of the devastating RNS. Therefore the aim of the present study was to examine the impact of dietary iron deficiency on cardiac mitochondrial microscopic morphology and cytochrome c release, NOS expression in both whole-cell extraction and mitochondria, and protein nitrotyrosine formation as an indicator of RNS. Several other related cardiac stress markers such as NADPH oxidase, caveolin-1 and RhoA, an important G-protein, which is closely associated with sarcomere rearrangement during cardiac remodelling [13], were determined in ventricular tissues from adult rats.


Animals and dietary feeding regime

The experimental procedures used in the present study were approved by the University of Wyoming Animal Use and Care Committee. In brief, 1-month-old male Sprague–Dawley rats were obtained from Charles River Laboratories and randomly divided into two groups fed either an iron-deficient diet (Diet #115109; Dyets Inc.) or a control diet (AIN-93G; Dyets Inc.) for 12 weeks. The iron-deficient diet was AIN-93G with reagent-grade salts (for low-iron purity) and no added iron. Rats were fed and given water (purified by reverse osmosis) ad libitum, kept on a 12 h light/dark cycle, weighed weekly, and maintained in accordance with the National Academy of Science’s Guide for the Care and Use of Laboratory Animals. BW (body weight) and organ weight were measured with a laboratory scales at the completion of the feeding period [6].

Morphometric measurement of hearts

The experimental animals were killed under anaesthesia and the hearts were cut open along the sagittal axis. Ventricular walls and chambers were examined ventrally. Morphometric measurements were performed using metric calipers and an 8 × Agfa magnifying loupe. Cardiac length and width, right and LV (left ventricular) free wall thickness, and LV major and lesser diameter were measured to the nearest 0.2 mm. The LV major length was the distance perpendicular from a line drawn joining cranial aspects of mitral and tricuspid valves to the distal aspect of the LV lumen. LV lesser diameter was the width of the LV lumen. LV chamber volume was calculated as (4/3)π×(LV major length/2)×(LV lesser diameter/2)2 [2].

Transmission electron microscopy

Tissue sections of the left ventricles of two rats selected randomly from each group were fixed with 2.5% glutaraldehyde/1.2% acrolein in fixative buffer [0.1 M cacodylate and 0.1 M sucrose (pH 7.4)], post-fixed with 1% osmium tetroxide, followed by 1% uranyl acetate, dehydrated through a graded series of ethanol concentrations, and embedded in LX112 resin (LADD Research Industries). Sections (1 μm thick) were cut on an ultramicrotome (RMC-MTXL), stained with 1% Toluidine Blue in 1% sodium borate, and viewed on an Axiophot light microscope (Carl Zeiss). Ultra-thin sections (approx. 50 nm) were cut on the ultramicrotome, stained with uranyl acetate, followed by lead citrate, and viewed on a 1200EX transmission Electron Microscope (Hitachi-7000) equipped with a 4K digital camera at 80 kV [14,15].

Analysis of mitochondrial cytochrome c release

The ventricles were minced and homogenized using a Polytron homogenizer in ice-cold MSE buffer {220 mM mannitol, 70 mM sucrose, 2 mM EGTA, 5 mM Mops (pH 7.4), 0.2% BSA and a protease inhibitor cocktail containing AEBSF [4-(2-aminoethyl)benzenesulphonyl fluoride], E-64, bestatin, leupeptin, aprotinin and EDTA (Sigma)}. The homogenates were centrifuged for 10 min at 600 g to remove unbroken tissue and nuclei, and the supernatants were centrifuged for 10 min at 3000 g to pellet mitochondria. The supernatants were centrifuged further for 30 min at 100000 g to obtain the cytosolic fraction. The mitochondrial pellet was dissolved in lysis buffer [20 mM Tris/HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton, 0.1% SDS and 1% protease inhibitor cocktail] and centrifuged at 10000 g for 30 min at 4 °C to produce a soluble protein fraction. A portion (50 μg) of the mitochondrial protein or cytosolic protein fraction was separated by SDS/PAGE [15% (w/v) acrylamide] and analysed by Western blot using anti-cytochrome c antibody (Upstate) [16].

Western blot analysis of eNOS, iNOS, p47phox subunit of NADPH oxidase, caveolin, RhoA and β-actin

Total protein (whole-cell extraction) was prepared as described previously [17]. In brief, tissue samples from heart ventricles were removed and homogenized in lysis buffer. Samples were then sonicated for 15 s and centrifuged at 12000 g for 20 min at 4 °C. The protein concentration of the supernatant was evaluated using Protein Assay Reagent (Bio-Rad Laboratories). Equal amounts (50 μg of protein/lane) of the protein from the whole-cell extraction, or mitochondria and prestained molecular-mass markers (Gibco-BRL), were separated on 7% (iNOS and eNOS), 10% (p47phox and RhoA) or 15% (caveolin-1 and β-actin) (w/v) polyacrylamide gels in a minigel apparatus (Mini-PROTEAN II; Bio-Rad Laboratories). Separated proteins were then transferred electrophoretically on to nitrocellulose membranes (0.2 μm pore size; Bio-Rad Laboratories). Membranes were incubated for 1 h in a blocking solution [TBS (Tris-buffered saline; 20 mM NaCl and 2mM Tris/HCl, pH 7.4) containing 5% (w/v) non-fat milk], washed briefly in TBS, and then incubated overnight at 4 °C with the appropriate dilution of antibodies: anti-p47phox (1:1000 dilution; monoclonal antibody kindly provided by Dr Mark T. Quinn, Montana State University, Bozeman, MT, U.S.A.), anti-eNOS (1:2000 dilution; BD Pharmigen), anti-iNOS (1:1000 dilution; Santa Cruz Biotechnology), anti-RhoA (1:1000 dilution; Upstate), anti-caveolin-1 (1:1000 dilution; Sigma) and β-actin (1:5000 dilution; Cell Signaling Technology) antibodies. After washing the blots to remove excess primary antibody binding, blots were incubated for 1 h with HRP (horseradish peroxidase)-conjugated secondary antibody (1:5000 dilution). Antibody binding was detected using enhanced chemiluminescence (Amersham Biosciences), and the resulting film was scanned and the intensity of immunoblot bands was detected with a calibrated densitometer (Model GS-800; Bio-Rad Laboratories).

Myocardial nitrotyrosine content

Nitrotyrosine content, a footprint of in vivo ONOO formation, was determined using an ELISA method. Myocardial tissue was homogenized in ice-cold PBS (1/10, w/v) using a PRO 200 homogenizer, followed by sonication with a dismembrator (medium intensity for 30 s; Fisher Scientific). Homogenates were centrifuged for 10 min at 12500 g at 4 °C. Supernatants were collected and protein concentrations were determined. Tissue samples from hearts (50 μg of protein) were applied on to disposable sterile ELISA plates (Corning Glassworks) and incubated overnight at 4 °C. The plate was then washed with 200 μl of PBS/0.1% Tween buffer, followed by incubation with heat-inactivated 10% (v/v) goat serum in PBS for 1 h at 37 °C to block non-specific binding. The primary antibody (mouse monoclonal antibody against nitrotyrosine; 1:2000 dilution; incubation for 2 h at 37 °C; Alexis Biochemicals) and secondary antibody (HRP-conjugated anti-mouse IgG; 1:2000 dilution; incubation for 1.5 h at 37 °C) were added, and the peroxidase reaction product was then generated using 2.2 mM O-phenylenediamine dihydrochloride solution (Abbott Diagnostics). Plates were incubated for 20 min in the dark at room temperature, and the reaction was stopped by addition of 50 μl of 1 M HCl to each well. Absorbance was measured at 450 nm with a Spectra Max 190 Microplate Spectrophotometer (Molecular Devices) [18].

Statistical analysis

Due to specific requirements of the measurement technique, it was not possible to use the same sample for all experimental indices. In this case, different rats were used for different parameters using randomized sample selection criteria. Data are expressed as means±S.E.M. Differences between variables were analysed by the non-parametric Mann–Whitney rank sum test (SPSS). The pattern of data distribution (normality) was assessed using Kolmogorov–Smirnov test (SAS software). All data in Table 2, with the exception of LV lesser diameter and LV chamber volume, were non-normally distributed. The data in Figures 3–6 that were non-normally distributed included control mitochondrial cytochrome c release, control whole-cell homogenate eNOS, mitochondrial eNOS and iNOS (except for control iNOS), p47phox subunit of NADPH oxidase, RhoA and caveolin-1 expression (except for p47phox in the iron-deficient group), and all data used for protein nitrotyrosine formation (except for the whole-cell formation in iron-deficient group). P<0.05 was considered statistically significant.


General features of experimental animals fed the control or iron-deficient diets

As shown in Table 1, rats consuming an iron-deficient diet were anaemic (as assessed by significantly reduced haematocrit values) compared with the control rats. Feeding rats the iron-deficient diet for 12 weeks did not significantly affect BW and liver weight or size of liver (organ/BW ratio) compared with the control diet. However, kidney and brain organ weights as well as brain size were significantly smaller in rats fed the iron-deficient diet compared with the control group. Kidney size (kidney weight/BW ratio) was comparable between the two dietary groups. As expected, absolute heart weight and heart size (heart/BW ratio) were both significantly larger in the iron-deficient group compared with the control group. As shown in Table 2, the heart length, heart width, and LV and right ventricular free wall thickness were all similar between the control and iron-deficient groups. However, following 12 weeks of feeding with the iron-deficient diet, LV major length, LV lesser diameter and LV chamber volume were all significantly increased compared with the control group, suggesting the presence of dilated cardiomyopathy.

Table 1General features of adult male rats fed the control or iron-deficient diets

Values are means±S.E.M. *P<0.05 compared with the control group.

Table 2Cardiac morphometry in rats fed the control or iron-deficient diets

Values are means±S.E.M. *P<0.05 compared with the control group.

Electron microscopic characteristics of hearts from rats fed control or iron-deficient diets

No ultrastructural abnormalities were observed in cardiac samples from ventricular samples of rats fed the control diet (Figures 1A and 1C). However, feeding rats an iron-deficient diet induced extensive focal damage in myocardial tissue sections (Figures 1B and 1D). This was indicated by cyto-architectural damage, including mitochondrial swelling, changes in shape and loss of integrity, significant disruption of sarcomeres and the arrangement of cardiac contractile filaments. However, the number of mitochondria did not differ in myocardial sections between the two groups (52.833±4.875 in controls compared with 49.714±4.144 in iron-deficient rats). Ultrastructural microscopy also revealed changes in sarcomeres in ventricular myocytes from rats fed the iron-deficient diet compared with those on the control diet. Figure 2 shows that LV myocytes from rats fed the iron-deficient diet displayed severely disrupted sarcomere and interfibrillar mitochondrial structure in comparison with the typically organized sarcomere and interfibrillar mitochondria in LV myocytes from rats fed the control diet.

Figure 1Transmission electron microscopic micrographs exhibiting ultrastructural changes and number of mitochondria in LV myocytes from rats fed with control or iron-deficient diet

Mitochondria from LV myocytes from rats fed the control diet (A) and iron-deficient diet (B). In (B), note that mitochondria are swollen with altered morphology. Magnification, ×15000. Higher-power magnification (×30000) of mitochondria from LV myocytes from rats fed the control (C) and iron-deficient (D) diets. In (D), note the degenerated mitochondria with membrane loss.

Figure 2Ultrastructural microscopic changes of sarcomere in ventricular myocytes from rats fed the control or iron-deficient diets

(A) LV myocytes from rats fed the control diet showing typically organized sarcomere structure and interfibrillar mitochondria. (B) LV myocytes from rats fed the iron-deficient diet exhibiting disrupted sarcomeres. Magnification, ×8000.

Effects of dietary iron-deficiency on cytochrome c release, protein expression of eNOS, iNOS, NADPH oxidase, RhoA, caveolin-1 and β-actin, and protein nitrotyrosine formation

To elucidate the possible contributing factors responsible for dietary iron-deficiency-induced cardiac enlargement, mitochondrial damage and sarcomere disruption, we examined mitochondrial cytochrome c release, a marker of mitochondrial damage, and several stress signalling molecules involved in RNS-mediated mitochondrial injury. Immunoblot analysis revealed that the mitochondrial cytochrome c release was significantly elevated (by approx. 70%) in the iron-deficient group compared with that from the control group (Figure 3). Expression of the housekeeping protein β-actin was similar between the two groups (Figure 3). As shown in Figure 4, the protein expression of eNOS and iNOS were significantly higher in whole-cell homogenates from the iron-deficient group. Protein expression of iNOS, but not eNOS, was significantly increased in the mitochondrial fractions from the iron-deficient group. Protein expression of the p47phox subunit of NADPH oxidase (Figure 5B), the membrane structural protein caveolin-1 (Figure 6A) and the small G-protein RhoA (Figure 6B) were all significantly increased in ventricular tissue from the iron-deficient group compared with the control group. Finally, as shown in Figure 5(A), protein nitrotyrosine formation, a key marker for RNS-induced protein injury, as detected by ELISA, was significantly increased in the mitochondrial, but not whole-cell, fraction from the iron-deficient group compared with the control group.

Figure 3Effect of dietary iron deficiency on cardiac cytochrome c release

Ventricular tissues from rats fed the control or iron-deficient diets were homogenized and separated by differential density centrifugation to yield membrane (mitochondria) and soluble (cytosol) fractions. Membrane and soluble proteins (50 μg) were separated by SDS/PAGE and analysed by Western blotting using an anti-cytochrome c antibody (Cyt C) or anti-β-actin antibody, a housekeeping protein used as an internal control. Quantification of protein expression (means±S.E.M.) is also shown; n=12. *P<0.05 compared with control group.

Figure 4Effect of dietary iron deficiency on cardiac protein expression of eNOS (A) and iNOS (B)

Representative blots probed with anti-eNOS or -iNOS antibodies in heart mitochondria and whole-cell extracts are shown. Quantification of protein expression (means±S.E.M.) is also shown; n=8–9 per group. *P<0.05 compared with control.

Figure 5Effect of dietary iron deficiency on cardiac protein nitrotyrosine formation (A) and expression of NADPH oxidase p47phox subunit (B)

(A) Nitrotyrosine formation was determined by ELISA. (B) Representative blot showing immunostaining with an anti-p47phox antibody in ventricular tissue. Quantification of protein expression (means±S.E.M.) is also shown; n=4 per group. *P<0.05 compared with control group.

Figure 6Effect of dietary iron deficiency on cardiac expression of caveolin-1 (A) and RhoA (B)

Representative blots showing immunostaining with anti-caveolin-1 and -RhoA antibodies. Cardiac proteins (20 and 50 μg respectively) were separated by SDS/PAGE [20% or 10% (w/v) acrylamide respectively]. Quantification of protein expression (means±S.E.M.) is also shown; n=8 (caveolin-1) or 12 (RhoA) per group. *P<0.05 compared with control group.


The results of the present study show that dietary iron deficiency is associated with cardiac hypertrophy, ultrastructural changes in mitochondria and sarcomeres, and increased release of cytochrome c from mitochondria into cytosol in hearts. Enhanced NOS expression and elevated protein nitrotyrosine formation were demonstrated for the first time in hearts from iron-deficient rats. In addition, increased expression of the p47phox subunit of NADPH oxidase in hearts from iron-deficient rats suggested an alternative mechanism of elevated nitrotyrosine formation through O2-mediated production of ONOO. Caveolin-1, a constituent protein of caveolae, regulates NO signalling by inhibiting eNOS [19]. Its expression was up-regulated in hearts from rats fed an iron-deficient diet, indicating multi-regulation of NO signalling in cardiac damage. Significantly enhanced levels of RhoA protein might suggest a contribution of small G-protein signalling to sarcomere disruption in hearts from iron-deficient rats.

A previous investigation [20] has suggested that altered myocardial function develops in iron-deficient rats with changes in ventricular contractility, maximal rates of contraction and relaxation, action potentials and L-type Ca2+ currents. In addition, the twitch duration was prolonged similar to that observed in diabetes and heart failure [20]. These electrophysiological changes may contribute to depressed cardiac function with iron deficiency. On the other hand, dietary iron deficiency has been shown to induce cardiac eccentric hypertrophy [2], further indicating the aetiology of heart dysfunction in anaemic (iron-deficient) patients. The morphological findings from our present study revealed that the major/minor ventricular diameters and ventricular volume were significantly enhanced in the iron-deficient group, consistent with cardiac hypertrophy (absolute heart weight and heart weight/BW ratio) observed in hearts from iron-deficient rats.

Mitochondria generate energy and heat by metabolizing oxygen and nutrients in addition to regulation of cellular metabolism and apoptosis [21]. Our results revealed an enhanced cytosolic fraction of cytochrome c associated with reduced mitochondrial cytochrome c levels in hearts from iron-deficient rats, indicating a ‘translocation’ of cytochrome c from mitochondria, which is a hallmark of mitochondrial injury and a key pre-apoptotic mitochondrial event. Our data also showed elevated mitochondrial nitrotyrosine formation in iron-deficient groups, indicating NO-related cardiac damage. NO controls oxygen supply and respiratory substrates to mitochondria and redistribution of heat generated by mitochondria [22]. However, NO itself could be cytotoxic, since it may inactivate mitochondrial respiratory chain enzymes and directly stimulate the mitochondrial pathway of apoptosis. Recent evidence suggests that nitrogen-derived stress is a unique form of cell injury under a wide variety of pathophysiological conditions [12]. Enhanced nitrotyrosine formation, a footprint of ONOO formation, has been demonstrated in a number of pathological conditions such as ischaemia/reperfusion injury [23]. Excessive NO produced from eNOS, or more frequently iNOS, may react with O2 to form ONOO, resulting in protein nitrotyrosine formation. Both eNOS and iNOS were found in mitochondria in cardiac myocytes [24,25]. Meanwhile, iNOS was induced in mitochondria providing a possible role of RNS in mitochondrial damage in iron deficiency. Although we did not measure local NOx (nitrite/nitrate), enhanced eNOS and iNOS expression observed in our present study supports the observation of enhanced protein nitrotyrosine formation due to elevated NOx levels [25a].

Our present study also revealed that iron deficiency may trigger the up-regulation of stress signalling molecules, including NADPH oxidase and RhoA. NADPH oxidase activation may directly promote generation of free radicals such as O2, leading to inactivation of NO and production of ONOO [26]. The up-regulation of NADPH oxidase under iron deficiency coincides with elevated eNOS, iNOS and protein nitrotyrosine formation in our present study. On the other hand, RhoA regulates cell morphology and contraction. Cardiac-specific overexpression of RhoA leads to prolongation of action potential duration and diminished ventricular contractility [27,28], suggesting a role of RhoA in impaired myocardial function in iron deficiency. RhoA is essential for the progression of cardiac hypertrophy through the regulation of the actin/myosin cytoskeleton. The effects of RhoA on cellular architecture may be mediated through Rho-dependent Ser/Thr protein kinases. RhoA activation of Rho kinase promotes phosphorylation of MLC (myosin light chain), although this increase in MLC phosphorylation regulates cytoskeletal organization [13]. Thus the data also suggest that up-regulated RhoA might contribute to aberrant sarcomere organization. One interesting finding from the present study is the elevated caveolin-1 expression in iron deficiency. Caveolin-1 acts through packing proteins in caveolae by protein–protein interactions, allowing finely tuned regulation of physiological responses [29]. Caveolin-1 usually suppresses growth and cell proliferation and is closely associated with apoptosis [3032]. Although the mechanism of action of elevated caveolin-1 expression is not clear at present, one would speculate that elevated expression of caveolin-1 during iron deficiency may sensitize myocardial cells to apoptotic stimuli via activation of apoptotic enzymes (such as caspase 3) [33]. Caevolin-1 has been suggested to participate in the regulation of eNOS protein function and may thus contribute to iron-deficiency-induced cardiac damage through an eNOS-dependent mechanism [34].

In a model of physiological stress as multifaceted as iron deficiency, it is difficult to distinguish between changes that result from lack of dietary iron, from anaemia or tissue hypoxia, or from compensatory physiological responses to anaemia. For example, iron deficiency resulted in a dilated eccentric cardiac hypertrophy, as demonstrated in the present study and elsewhere [2]. This morphological alteration was found whether anaemia was severe [2] or moderate, as in our present study (Table 1). With dietary copper deficiency, a concentric hypertrophic pattern (increased ventricular wall thickness in the absence of increased chamber size) was found without or with anaemia [2,35]. This might suggest that the iron-deficiency-induced dilated hypertrophy pattern is secondary to the physiological stimuli related to anaemia (whether or not that anaemia is severe), whereas copper-deficiency-induced hypertrophy [35] is unrelated to anaemia. Increased venous return, decreased peripheral resistance, increased inotropic response to noradrenaline (norepinephrine) and vascular remodelling to a larger arterial diameter have all been demonstrated with iron-deficiency-induced anaemia [57]; a chronic hypersympathetic state coupled with the haemodynamic changes has been suggested to be essential for cardiac hypertrophy [6]. In contrast, copper-deficiency-induced hypertrophy has been attributed to mitochondrial proliferation [36].

Similarly, the mitochondrial changes reported in the present study appear to have significant differences compared with the copper-deficiency model as a direct result of disparate mineral deficiencies. For example, copper deficiency induces an increase in cardiac mitochondrial number, which may allow normal levels of oxidative phosphorylation [37]. We found no increase in mitochondrial number with iron deficiency; furthermore, it would appear unlikely that oxidative phosphorylation is unchanged with iron deficiency given our findings of ultrastructural damage to mitochondria, the translocation of cytochrome c from mitochondria to cytoplasm and an unchanged mitochondrial number. Copper deficiency has also been shown [38] to reduce the number of subunits of cytochrome c oxidase that are transcribed in the nucleus, but not mitochondria. However, this is not true with iron deficiency [38]. Thus, although electron transport Complex IV requires both iron and copper to function properly and both deficiencies result in cardiac hypertrophy [2,35,36], there appear to be significant differences in cardiac mitochondrial alterations that are best explained as direct responses to mineral deficiency.

Experimental limitations of the present study

One major limitation of the present investigation is that our conclusions from this ‘uncontrolled comparison’ study are essentially based upon observed differences between groups, rather than any cross-sectional associations between parameters within the same group. Therefore any causal relationship between reactive oxygen species and cardiac damage during iron deficiency cannot be drawn in a convincing manner at the present stage. In addition, since it was impossible to use the same sample for all experimental indices, use of different rats for different parameters reduces the potential usefulness of examining associations between various parameters. This also contributed to the relatively small sample sizes (i.e. different rats were, in some instances, used for different parameters due to requirement of the measuring technique). Nevertheless, samples were randomly chosen for any parameter tested (if not all rat tissue samples were available for that given parameter).

In summary, the findings of the present study have shown that dietary iron deficiency and/or anaemia may trigger alterations in cardiac morphology, including enlarged (dilated) ventricles, ultrastructural aberrations in mitochondria and sarcomeres, elevated release of cytochrome c from mitochondria, elevated protein nitrotyrosine formation, and higher levels of eNOS, iNOS, NADPH oxidase, caveolin-1 and RhoA. These data suggest the presence of RNS and cardiac damage during iron deficiency, and may enhance our understanding of the mechanism and possible therapeutic regime for dietary iron-deficiency-related cardiac dysfunction.


We thank Dr Xinrong Qiang from the University of Wyoming for her assistance with the statistical analysis. We also thank Dr Zhaojie Zhang from the University of Wyoming Microscopy Facility and Ms Resha Ball from Western Wyoming College for their assistance during this study. This work was supported, in part, by grants from the National Institute of Health (NCRR BRIN P20 RR15640, NCRR COBRE P20 RR15640 and NIH R03 AG21324-01), and the American Heart Association Pacific Mountain Affiliate (0355521Z).

Abbreviations: BW, body weight; HRP, horseradish peroxidase; LV, left ventricular; MLC, myosin light chain; NO, nitric oxide; NOS, NO synthase; eNOS, endothelial NOS; iNOS, inducible NOS; NOx, nitrite/nitrate; O2−, superoxide anion; ONOO−, peroxynitrite; RNS, reactive nitrogen species; TBS, Tris-buffered saline

Why is iron in your products? We hear today 
that iron is dangerous especially for men.

Iron is an essential nutrient. It is on the list of required nutrients. While an excess of any nutrient is not good, iron is still an essential part of good nutrition. Excess iron may be caused by a variety of factors, many of which may be discovered by a qualified medical doctor. If there is no apparent reason for elevated iron it may be an imbalance between minerals. Our products will re-balance the body so that it may more easily find it’s own equilibrium. The Majestic Earth products only provide a part of the minimum daily intake of iron.

Iron: What is it?
Iron is an essential mineral and an important component of proteins involved in oxygen transport and metabolism (1,2). Almost two-thirds of the iron in your body is found in hemoglobin, the protein in red blood cells that carries oxygen to your body’s tissues. Smaller amounts of iron are found in myoglobin, a protein that helps supply oxygen to muscle, and in enzymes that assist biochemical reactions in cells.

About 15 percent of your body’s iron is stored for future needs and mobilized when dietary intake is inadequate. The remainder is in your body’s tissues as part of proteins that help your body function. Adult men and post-menopausal women lose very little iron except through bleeding.

Women with heavy monthly periods can lose a significant amount of iron. Your body usually maintains normal iron status by controlling the amount of iron absorbed from food.

What is the Recommended Dietary Allowance for Iron
The Recommended Dietary Allowance (RDA) is the daily dietary intake level that is sufficient to meet the nutrient requirements of nearly all (97-98%) healthy individuals in each life-stage and gender group (1).

The 2001 RDAs for iron (in milligrams) for infants ages 7 to 12 months, children and adults are:

Age  Infants, Children  Males  Females Pregnancy  Lactation
7 to 12 months 11 mg
1 to 3 years 7 mg
4 to 8 years 1o mg
9 to 13 years 8 mg 8 mg
14-18 years 11 mg 15 mg 27 mg 10 mg
19-50 years 8 mg 18 mg 27 mg 9 mg
51+ years 8 mg 8 mg

When can iron deficiency occur?
The World Health Organization considers iron deficiency the number one nutritional disorder in the world. It affects more than 30% of the world’s population.

When your need for iron increases or a loss of iron from bleeding exceeds your dietary iron intake, a negative iron balance may occur. Initially this results in iron depletion, in which the storage form of iron is decreased while blood hemoglobin level remains normal. Iron deficiency occurs when blood and storage levels of iron are low, and the blood hemoglobin level falls below normal .

Iron deficiency anemia may result from a low dietary intake, inadequate intestinal absorption, excessive blood loss, and/or increased needs. Women of childbearing age, pregnant women, older infants and toddlers, and teenage girls are at greatest risk of developing iron deficiency anemia because they have the greatest needs .

Individuals with renal failure, especially those receiving dialysis, are at high risk for developing iron deficiency anemia. This is because their kidneys cannot create enough erythropoietin, a hormone needed to make red blood cells. Iron and erythropoietin can also be lost with blood during dialysis, which can result in an iron deficiency. Extra iron and erythropoietin are usually needed to help prevent iron deficiency in these individuals .

Iron deficiency could also be caused by low vitamin A status. Vitamin A helps to mobilize iron from its storage sites, so a deficiency of vitamin A limits the body’s ability to use stored iron. This results in an “apparent” iron deficiency because hemoglobin levels are low, even though the body can maintain normal amounts of stored iron . While uncommon in the U.S., this problem is seen in developing countries where vitamin A deficiency often occurs.

The anemia that may occur with inflammatory disease differs from iron deficiency anemia. It occurs in people who have chronic infectious, inflammatory, or malignant disorders . It is not associated with a shortage of dietary iron, and may not respond to iron supplementation. A physician should manage anemia associated with an inflammatory disorder.

Signs of iron deficiency anemia include feeling tired and weak, decreased work and school performance, slow cognitive and social development during childhood, difficulty maintaining body temperature, and decreased immune function, which may decrease resistance to infection . During pregnancy, iron deficiency is associated with increased risk of premature deliveries, giving birth to infants with low birth weight, and maternal complications .

Iron and Heart Disease.
Several observations have led researchers to examine the association between high iron stores and coronary heart disease. It appears that rates of heart disease among women increase when monthly periods stop, a time when levels of stored iron increase. Also, some researchers have suggested that lower rates of heart disease among people living in developing countries may be due to low meat (and iron) intake, high fiber diets that inhibit iron absorption, and gastrointestinal (GI) parasite concentrations that result in gastrointestinal blood (and iron) loss, all of which contribute to low iron stores in this population. In addition, a 1980s study of Finnish men linked high iron stores with increased risk of heart attacks. However, not all studies have supported this relationship , including a 1999 review of 12 studies that failed to show a strong association . It is also true that older women have a greater prevalence of traditional cardiovascular disease risk factors such as high blood pressure and elevated blood cholesterol. Currently, available data do not provide convincing support for an association between high body iron stores and increased risk for coronary heart disease .

What is the health risk of too much iron?

Iron has a moderate to high potential for toxicity because very little iron is excreted from the body. Thus, iron can accumulate in body tissues and organs when normal storage sites are full.

In children, acute toxicity can occur from overdoses of medicinal iron. Ingestion of as few as five or six high-potency tablets can provide amounts of iron that can be fatal to a child of 22 pounds. Consuming 1 to 3 grams of iron can be fatal to children under six and lower doses can cause severe symptoms such as vomiting and diarrhea (64). It is important to keep iron supplements tightly capped and away from children’s reach. Any time excessive iron intake is suspected, immediately call your physician or Poison Control Center, or visit your local emergency room. In adults high intakes of iron supplements are associated with constipation, nausea, vomiting, and diarrhea, especially when the supplements are taken on an empty stomach (1).

In 2001, the Institute of Medicine set a tolerable upper intake level (UL) of 40 mg per day for infants and children through age 13 and 45 mg per day for adolescents ages 14 to 18 years and adults 19 years of age and older (1). The upper limit does not apply to individuals who receive iron under medical supervision. There may be times when a medical doctor prescribes an intake higher than the upper limit, such as when individuals with iron deficiency anemia need higher doses of iron until their iron stores return to normal.

National Institutes of Health (NIH), Bethesda, MD, in conjunction with the Office of Dietary Supplements (ODS) in the Office of the Director of NIH.

Joel Wallach