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An essential cell-autonomous role for hepcidin in cardiac iron homeostasis

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Cite this article as: eLife 2016;5:e19804 doi: 10.7554/eLife.19804

Abstract

Hepcidin is the master regulator of systemic iron homeostasis. Derived primarily from the liver, it inhibits the iron exporter ferroportin in the gut and spleen, the sites of iron absorption and recycling respectively. Recently, we demonstrated that ferroportin is also found in cardiomyocytes, and that its cardiac-specific deletion leads to fatal cardiac iron overload. Hepcidin is also expressed in cardiomyocytes, where its function remains unknown. To define the function of cardiomyocyte hepcidin, we generated mice with cardiomyocyte-specific deletion of hepcidin, or knock-in of hepcidin-resistant ferroportin. We find that while both models maintain normal systemic iron homeostasis, they nonetheless develop fatal contractile and metabolic dysfunction as a consequence of cardiomyocyte iron deficiency. These findings are the first demonstration of a cell-autonomous role for hepcidin in iron homeostasis. They raise the possibility that such function may also be important in other tissues that express both hepcidin and ferroportin, such as the kidney and the brain.

https://doi.org/10.7554/eLife.19804.001

eLife digest

Many proteins inside cells require iron to work properly, and so this mineral is an essential part of the diets of most mammals. However, because too much iron in the body is also bad for health, mammals possess several proteins whose role is to maintain the balance of iron. Two proteins in particular, called hepcidin and ferroportin, are thought to be important in this process. Some ferroportin is found in the cells that line the gut (where iron is absorbed into the body) and is required to release this iron into the bloodstream. It is also found in the spleen, which is where iron is removed from old red blood cells so that it can be recycled. The liver produces hepcidin to control when ferroportin is active in the gut and spleen.

Both hepcidin and ferroportin are also found in heart cells. In 2015, a study reported that that heart ferroportin plays an important role in heart activity. However, it was not clear what role hepcidin plays in this organ.

Now, Lakhal-Littleton et al. – including many of the researchers from the previous work – have genetically engineered mice such that they specifically lacked heart hepcidin, or had a version of ferroportin in their heart that does not respond to hepcidin. The experiments show that these changes caused fatal heart failure in the mice because ferroportin releases iron from heart cells in an uncontrolled manner. Lakhal-Littleton et al. were able to prevent heart failure by injecting the animals with iron directly into the bloodstream. These findings show that hepcidin produced outside the liver has a role in controlling the levels of iron in the body’s organs.

Other organs such as the brain, kidney and placenta all have their own forms of hepcidin and ferroportin; further work could investigate the roles of these proteins. Finally, another challenge for the future will be to test whether new drugs that are being developed to block or mimic hepcidin from the liver have the potential to treat heart conditions in humans.

https://doi.org/10.7554/eLife.19804.002

Introduction

As a constituent of hemoproteins, iron-sulphur proteins and other functional groups, iron is essential for cellular functions. Conversely, excess iron participates in cytotoxic Fenton-type chemical reactions. Thus, both iron deficiency and iron overload are detrimental to the cell. Therefore, the healthy functioning of tissues requires tight control of intracellular iron levels. These in turn are dependent both on cellular homeostatic pathways controlling iron uptake, usage, and storage, and on systemic pathways controlling iron levels in the plasma. In mammals, cellular iron homeostasis is controlled by the Iron Regulatory Proteins IRPs. Intracellular iron levels control the degradation of IRP2 and the conformational switch that confers the RNA-binding function of IRP1. IRPs in turn control the levels of iron uptake proteins such as transferrin receptor 1 (TfR1) and divalent metal transporter (DMT1), and the iron storage protein ferritin (Rouault and Klausner, 1997; Rouault, 2006). Systemic iron homeostasis is controlled by the hepcidin/ferroportin axis at the sites of iron entry into the circulation. Ferroportin (FPN), which is encoded by the Solute Carrier Family 40 Member 1 (Slc40a1), is the only known mammalian iron export protein and mediates iron release into the circulation from duodenal enterocytes, splenic reticuloendothelial macrophages and hepatocytes, the sites of iron absorption, recycling and storage respectively (Ganz, 2005; Donovan et al., 2005). FPN-mediated iron release is antagonized by the hormone hepcidin, also known as hepcidin antimicrobial peptide (HAMP). Produced primarily in the liver, hepcidin binds to and induces internalization of FPN, thereby limiting iron release into the circulation and its availability to peripheral tissues (Nemeth et al., 2004; Qiao et al., 2012). The importance of the HAMP/FPN axis is illustrated by diseases of systemic iron overload such as hemochromatosis and β-thalassaemia, where hepcidin production is impaired (Camaschella, 2005; Musallam et al., 2012), and in anaemia of chronic disease where hepcidin production is inappropriately elevated (De Falco et al., 2013; Nemeth and Ganz, 2014).

Other than the liver, hepcidin is also found in tissues with no recognized role in systemic iron homeostasis, including the heart (Merle et al., 2007), the brain (McCarthy and Kosman, 2014), the kidney (Kulaksiz, 2005) and the placenta (Evans et al., 2011). The function of this extra-hepatic hepcidin remains unknown, but one hypothesis is that it is involved in local iron control. Relevant to this hypothesis are our recent findings that FPN is also present in the heart, that it is essential for cardiomyocyte iron homeostasis and that its cardiomyocyte-specific deletion leads to fatal cardiac iron overload in mice (Lakhal-Littleton et al., 2015). Therefore, we hypothesised that cardiac HAMP regulates cardiac FPN, and that such regulation is important for local iron control and for cardiac function.

To test this hypothesis, we generated two novel mouse models; the first with a cardiomyocyte-specific deletion of the Hamp gene, and the second, with cardiomyocyte-specific knock-in of Slc40a1 C326Y, that encodes a FPN with intact iron export function but impaired HAMP binding (Schimanski et al., 2005; Drakesmith et al., 2005). We studied cardiac function and iron homeostasis longitudinally in both models and report that both develop fatal cardiac dysfunction and metabolic derangement as a consequence of cardiomyocyte iron deficiency. This occurs against a background of otherwise normal systemic iron homeostasis. Both cardiac dysfunction and metabolic derangement are prevented by intravenous iron supplementation.

Our findings demonstrate that, at least in the cardiomyocyte, endogenously-derived HAMP plays an essential role in cellular, rather than systemic iron homeostasis. It does this through the autocrine regulation of cardiomyocyte FPN. Disruption of this cardiac HAMP/FPN leads to fatal cardiac dysfunction.

Currently, there is considerable clinical interest in strategies that target the HAMP/FPN axis for the treatment of systemic iron overload and iron deficiency. Our findings suggest that these strategies may additionally alter cardiac iron homeostasis and function. Other than the heart, both HAMP and FPN are also found in the brain, kidney and placenta (McCarthy and Kosman, 2014; Kulaksiz, 2005; Evans et al., 2011; Rouault, 2013; Moos and Rosengren Nielsen, 2006; Bastin et al., 2006; Wolff et al., 2011). A pertinent question is the extent to which our findings in the heart extend to those tissues.

Results

Hamp expression and regulation in the heart

Hamp mRNA levels were approximately 30 fold lower in the adult mouse heart than in the liver (Figure 1A). Next, we examined the regulation of cardiac Hamp mRNA and HAMP protein in response to dietary iron manipulation, having first established that this dietary manipulation altered cardiac and liver iron levels (Figure 1—figure supplement 1). In both tissues, Hamp mRNA levels were decreased by the iron-deficient diet (Fe 2–5 ppm) and increased by the iron-loaded diet (Fe 5000 ppm) (Figure 1A). At the protein level, while changes in hepatic HAMP protein mirrored changes in its mRNA levels, cardiac HAMP protein was increased by the iron-deficient diet and unaffected by the iron-loaded diet (Figure 1B).

Figure 1 with 6 supplements see all
Hepcidin expression and regulation in the heart.

(A) Relative Hamp mRNA expression in heart and liver of adult C57BL/6 mice, under control conditions and after provision of low or high iron diets. *p=0.047, 0.001 respectively relative to control hearts, †p = 0.006, 0.019 respectively relative to control livers. (B) Corresponding immunohistochemical staining for HAMP in heart and liver. (C) Relative Hamp mRNA expression in primary adult mouse cardiomyocytes cultured under control conditions or in presence of FAC or DFO. *p=0.023, 0.001 and 0.014 respectively relative to control. †p = 0.024, 0.037, 0.016 and 0.037 respectively relative to control at the same timepoint. (D) Corresponding HAMP protein levels in supernatants of primary cardiomyocytes. DFO treatment was carried alone (DFO) or presence of Furin inhibitor (DFO+CMK). *p=0.002, 0.020, 0.028, 0.014, 0.015 respectively relative to control at the same timepoint. (E) Relative Hamp expression in heart and liver of 3 month old Hampfl/fl and Hampfl/fl;Myh6.Cre+ mice. *p=0.018 relative to cardiac Hamp in Hampfl/fl controls. (F) Corresponding immunohistochemical staining for HAMP in heart and liver. All values are plotted as mean ± SEM. Scale bar = 20 µm. n = 3 per group unless otherwise stated.

https://doi.org/10.7554/eLife.19804.003

To explore further the regulation of cardiac hepcidin by iron, we isolated primary adult cardiomyocytes from mice, then carried out a timecourse treatment with the iron chelator desferroxamine DFO, or with ferric citrate FAC. Under control conditions, Hamp mRNA was upregulated following addition of fresh cardiomyocyte culture medium (cardiomyocytes are cultured for 2 hr in serum-free medium prior to this). Relative to control cardiomyocytes at the respective timepoint, Hamp mRNA was increased by FAC from 4 hr of treatment, and decreased by DFO at 4, 8 and 16 hr of treatment (Figure 1C). HAMP protein, measured by ELISA in supernatants was also increased following addition of fresh medium. Relative to control cardiomyocytes at the respective timepoint, HAMP protein in supernatants was increased by DFO as early as 2 hr, but remained unchanged by FAC at all timepoints (Figure 1D). Thus, the direction of response of the Hamp mRNA and HAMP protein to iron levels in vitro mirrored the responses seen in vivo.

Next, we aimed to understand the mechanisms underlying increased HAMP secretion in DFO-treated cardiomyocytes. To this end, we investigated the role of the prohormone convertase Furin, which in hepatocytes, is required for cleavage of the propeptide and the release of the mature HAMP peptide (Valore and Ganz, 2008). Furin expression has been reported in the heart (Beaubien et al., 1995; Ichiki et al., 2013), and we also found that its expression was upregulated in the hearts of mice provided an iron-deficient diet and in cardiomyocytes treated with DFO (Figure 1—figure supplement 2). When we measured HAMP in supernatants of cardiomyocytes treated with the Furin inhibitor decanoyl-Arg-Val-Lys-Arg-chloromethylketone (CMK), we found no increase in HAMP release following DFO treatment (Figure 1D). We confirmed that Hamp mRNA levels in cardiomyocytes were not altered by CMK treatment (Figure 1—figure supplement 3). Together, these results indicate that Furin upregulation mediates increased HAMP secretion from iron-deficient cardiomyocytes.

Having established that hepcidin is found in cardiomyocytes, we then aimed to define its function. To this end, we generated cardiomyocyte-specific Hamp knockout mice Hampfl/fl;Myh6.Cre+ by crossing in-house conditional Hamp floxed (fl) mice with mice transgenic for Cre recombinase under the control of cardiomyocyte-specific promoter Myosin Heavy Chain 6 (Myh6.Cre+). Hamp mRNA (Figure 1E) and HAMP protein (Figure 1F) in the hearts of Hampfl/fl;Myh6.Cre+ mice were significantly reduced compared to the hearts of Hampfl/fl controls. Furthermore, compared to Hampfl/fl cardiomyocytes, levels of HAMP protein in the supernatants of cardiomyocytes from Hampfl/fl;Myh6.Cre+ mice were either greatly reduced or undetectable, both under baseline conditions and following treatment with DFO or FAC (Figure 1—figure supplement 4). Near complete ablation of the cardiac Hamp mRNA and HAMP protein in Hampfl/fl;Myh6.Cre+ mice confirmed that cardiomyocytes were the primary site of hepcidin expression in the heart. Liver Hamp mRNA (Figure 1E) and HAMP protein (Figure 1F) were not different between Hampfl/fl;Myh6.Cre+ and Hampfl/fl controls, consistent with the cardiac-specific nature of Hamp gene deletion.

Also consistent with this cardiac-specific deletion, Hampfl/fl;Myh6.Cre+ mice had normal levels of liver iron stores and circulating markers of iron homeostasis when compared to Hampfl/fl controls, demonstrating that loss of cardiac hepcidin did not affect systemic iron homeostasis. In addition, circulating HAMP levels were not reduced in the serum of Hampfl/fl;Myh6.Cre+ mice, suggesting that cardiac hepcidin does not contribute significantly to circulating HAMP levels (Table 1).

Table 1

Indices of systemic iron in six month old Hampfl/fl and Hampfl/fl;Myh6.Cre+ mice.

n = 6 per group. All values are shown as mean ± SEM.

https://doi.org/10.7554/eLife.19804.010

Hampfl/fl

Hampfl/fl;Myh6.Cre+

liver total elemental iron (ng/mg tissue)

96.3 ± 12.2

88.7 ± 19.2

liver ferritin (µg/mg total protein)

0.65 ± 0.04

0.64 ± 0.05

serum iron (µmol/L)

28.60 ± 7.20

31.50 ± 8.40

serum ferritin (mg/L)

1.81 ± 0.04

1.88 ± 0.29

hemoglobin (g/L)

122.7 ± 11.5

116.0 ± 11.9

serum hepcidin (µg/L)

23.5 ± 7.6

23.9 ± 10.40

Fatal cardiac abnormalities in Hampfl/fl;Myh6.Cre+ mice

To determine the effects of loss of cardiac hepcidin, we first assessed the cumulative survival of Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls over a period of 52 weeks. Significantly greater mortality was observed amongst Hampfl/fl;Myh6.Cre+ mice, with only 29% of animals surviving to 52 weeks, compared with 90% of Hampfl/fl controls. The median survival of Hampfl/fl;Myh6.Cre+ mice was 28 weeks, whereas the majority of Hampfl/fl controls were still alive at 52 weeks (Figure 2A).

Fatal cardiac abnormalities in Hampfl/fl;Myh6.Cre+ mice.

(A) Cumulative survival of Hampfl/fl;Myh6.Cre+ mice (n = 50) and Hampfl/fl littermate controls (n = 47) over 52 weeks. (B) Representative H and E longitudinal heart sections from a six month old Hampfl/fl;Myh6.Cre+ mouse and Hampfl/fl littermate control. (C) Representative WGA cardiac staining from a six month old Hampfl/fl;Myh6.Cre+ mouse and Hampfl/fl littermate control. (D) Quantitation of cardiomyocyte size based on WGA staining in six month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. *p=0.001 relative to Hampfl/fl littermate controls. (E) Relative expression of the hypertrophic gene markers Myh7 and Nppb in hearts of 6 month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. *p=0.001, 0.047 for the respective gene relative to Hampfl/fl littermate controls. (F) Representative cardiac in-situ TUNEL staining from a six month old Hampfl/fl;Myh6.Cre+ mouse and Hampfl/fl littermate control. (G) Quantitation of percentage of apoptotic cardiomyocytes based on in-situ TUNEL staining in six month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls *p=0.001 relative to Hampfl/fl littermate controls. (H) Representative midventricular Cine MR images of hearts from Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl controls at 3, 6 and 9 months of age. (IK) Cine MRI measurements of LV lumen, at end-systole (LVES), end-diastole (LVED), and of ejection fraction (LVEF) in Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls at three months (n = 8 per group), six months (n = 11 per group, *p=0.043 for LVES, 0.047 for LVED and 0.020 for LVEF) and nine months (n = 5 per group, *p=0.044 for LVES, 0.042 for LVED and 0.034 for LVEF). p values are relative to Hampfl/fl controls of the respective age. All Values are plotted as mean ± SEM. n = 3 per group unless otherwise stated. Scale bar = 50 µm.

https://doi.org/10.7554/eLife.19804.011

Six-month old mice were sacrificed for assessment of cardiac morphology, which showed gross enlargement of the left ventricle (LV) in Hampfl/fl;Myh6.Cre+ hearts compared to Hampfl/fl controls (Figure 2B). Assessment of cardiomyocyte size by wheat germ agglutinin (WGA) staining confirmed that Hampfl/fl;Myh6.Cre+ cardiomyocytes were hypertrophied (Figure 2C–D). This was accompanied by upregulation of expression of hypertrophic gene markers myosin heavy chain (Myh7) and natriuretic peptide precursor (Nppb) (Figure 2E). TUNEL staining for in-situ detection of cell death also showed significantly greater apoptosis in the hearts of Hampfl/fl;Myh6.Cre+ mice than in Hampfl/fl controls (Figure 2F–G).

To characterise further the phenotype caused by loss of cardiac hepcidin, we used cine MRI in anaesthetised Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls at 3, 6 and 9 months of age. Mid-ventricular cine MR images showed no differences between the two genotypes at 3 months of age. At 6 and 9 months of age, cine MR images showed marked enlargement of the LV in Hampfl/fl;Myh6.Cre+ mice compared to Hampfl/fl controls (Figure 2H). Formal quantitation of cardiac parameters by cine MRI confirmed enlargement of the LV lumen in Hampfl/fl;Myh6.Cre+ mice, both at end-systole (LVES) (Figure 2I) and at end-diastole (LVED) (Figure 2J), accompanied by a decrease in LV ejection fraction (LVEF) from 62% to 42% (Figure 2K). Other parameters of cardiac performance were not significantly altered between mice from the two genotypes (Table 2). Taken together, histological examination of the hearts and cine MRI studies indicated that Hampfl/fl;Myh6.Cre+ mice developed fatal LV dysfunction with reduced LVEF.

Table 2

Non-LV parameters of cardiac function are not altered between Hampfl/fl and Hampfl/fl;;Myh6.Cre+ mice.

Cine MRI measurements of cardiac function in Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl controls at three months (n = 8 per group), six months (n = 11 per group) and nine months (n = 5 per group) of age. Values are shown as mean ± SEM.

https://doi.org/10.7554/eLife.19804.013

 

3 months

6 months

9 months

Hamp fl/fl

Hamp fl/fl;

Myh6.Cre+

Hamp fl/fl

Hamp fl/fl;

Myh6.Cre+

Hamp fl/fl

Hamp fl/fl;

Myh6.Cre+

Average mass (mg)

70.51 ± 7.47

70.38 ± 5.33

72.59 ± 5.38

82.54 ± 11.32

78.30 ± 4.92

83.07 ± 5.44

RVED lumen (µl)

31.37 ± 2.55

26.81 ± 2.30

33.12 ± 3.19

30.63 ± 2.35

38.39 ± 3.88

39.95 ± 3.00

RVES lumen (µl)

7.43 ± 0.79

5.23 ± 0.67

8.70 ± 1.22

8.34 ± 1.06

13.36 ± 2.21

15.51 ± 2.47

RVEF (%)

76.42 ± 1.32

79.88 ± 2.80

73.66 ± 2.50

73.42 ± 2.22

65.20 ± 4.36

61.95 ± 3.53

Stroke volume (µl)

25.47 ± 1.99

22.73 ± 2.11

25.70 ± 2.70

23.76 ± 1.85

29.79 ± 2.30

29.84 ± 1.59

Cardiac output (ml/min)

10.34 ± 1.06

9.62 ± 0.90

10.72 ± 1.07

10.53 ± 1.05

11.46 ± 1.15

11.99 ± 1.38

Heart Rate (bpm)

404.07 ± 18.69

426.90 ± 17.94

419.64 ± 19.05

436.14 ± 14.30

384.84 ± 28.55

400.97 ± 41.20

Heart/body weight ratio x1000

2.80 ± 0.18

3.09 ± 0.26

2.51 ± 0.23

2.98 ± 0.37

2.36 ± 0.28

2.88 ± 0.27

Such changes in cardiac performance could not be attributed to Cre recombinase toxicity in the heart as we have previously shown that Myh6.Cre+ mice have normal cardiac function compared to wild type littermate controls (Lakhal-Littleton et al., 2015).

The role of cardiomyocyte FPN in the phenotype of Hampfl/fl;Myh6.Cre+ mice

We examined FPN protein in the hearts of Hampfl/fl;Myh6.Cre+ mice, and found that FPN protein was markedly upregulated compared to Hampfl/fl controls (Figure 3A), consistent with the idea that loss of cardiac HAMP was acting through upregulation of cardiomyocyte FPN. In order to test whether cardiac dysfunction arose from upregulation of cardiomyocyte FPN, we engineered mice where the Slc40a1 gene harbours a conditional cardiac-specific C326Y point mutation, which confers HAMP-resistance while conserving the iron export function of FPN (Schimanski et al., 2005; Drakesmith et al., 2005). We confirmed that the Slc40a1 C326Y fl allele produced the C326Y transcript specifically in the heart (Figure 3—figure supplement 1) and that Slc40a1 C326Yfl/fl;Myh6.Cre+ mice did not exhibit changes in systemic iron indices (Table 3). As seen in Hampfl/fl;Myh6.Cre+ mice, cardiomyocyte FPN was indeed upregulated in Slc40a1 C326Yfl/fl;Myh6.Cre+ mice (Figure 3B).

Figure 3 with 2 supplements see all
The role of cardiomyocyte FPN.

(AB) Immunohistochemical staining for FPN in the hearts of three month old Hampfl/fl;Myh6.Cre+, Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and respective controls. (C) Cumulative survival of Slc40a1 C326Yfl/fl;Myh6.Cre+ mice (n = 36) and Slc40a1 C326Yfl/fl littermate controls (n = 31) over 52 weeks. (D) Representative H and E-stained longitudinal heart sections from a six month old Slc40a1 C326Yfl/fl;Myh6.Cre+ mouse and a Slc40a1 C326Yfl/fl control. (E) Representative WGA cardiac staining from a six month old Slc40a1 C326Yfl/fl;Myh6.Cre+ mouse and Slc40a1 C326Yfl/fl control. (F) Quantitation of cardiomyocyte size based on WGA staining. n = 3 per group. *p=0.001 relative to Slc40a1 C326Yfl/fl controls. (G) Relative expression of Myh7 and Nppb in hearts of 6 month old Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and Slc40a1 C326Yfl/fl controls. n = 3 per group. *p=0.032, 0.044 for the respective gene relative to Slc40a1 C326Yfl/fl controls. (H) Representative cardiac TUNEL staining from a six month old Slc40a1 C326Yfl/fl;Myh6.Cre+ mouse and Slc40a1 C326Yfl/fl control. (I) Quantitation of percentage of apoptotic cardiomyocytes based on TUNEL staining, n = 3 per group. *p=0.0003 relative to Slc40a1 C326Yfl/fl controls. (JL) Cine MRI measurements of LVES, LVED and LVEF in Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and Slc40a1 C326Yfl/fl controls at three months (n = 6 per group), six months (n = 6 per group, *p=0.003 for LVES, 0.043 for LVED and 0.001 for LVEF) and nine months (n = 5 per group, *p=0.033 for LVES, 0.047 for LVED and 0.023 for LVEF). P values are relative to Slc40a1 C326Yfl/fl controls of the same age. (M) Percentage Fe55 efflux in cardiomyocytes from Hampfl/fl;Myh6.Cre+ mice, Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and respective controls, in presence or absence of HAMP peptide. *p=0.018, 0.006 and 0.007 respectively (N) Elemental iron levels in cardiomyocyte fractions (CF) from the hearts of Hampfl/fl;Myh6.Cre+ mice, Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and their respective controls. n = 4 per group. *p=0.032, 0.044, 0.047 and 0.031 respectively. (OP) Relative TfR1 (*p=0.038, 0.001) and Fpn (*p=0.039, 0.047) expression in hearts of 3 month old Hampfl/fl;Myh6.Cre+ mice, Slc40a1 C326Yfl/fl;Myh6.Cre+ mice and their respective controls. All values are plotted as mean ± SEM. Scale bar = 50 µm.

https://doi.org/10.7554/eLife.19804.014
Table 3

Characterisation of Slc40a1 C326Yfl/fl;Myh6.Cre+ mice.

Indices of iron status in Slc40a1 C326Yfl/fl and Slc40a1 C326Yfl/fl;Myh6.Cre+ mice at six months of age (n = 4 per group). Values are shown as mean ± SEM.

https://doi.org/10.7554/eLife.19804.018

 

Slc40a1 C326Yfl/fl

Slc40a1 C326Y fl/fl;

Myh6.Cre+

liver total elemental iron (ng/mg tissue)

92.77 ± 21.30

84.00 ± 26.00

liver ferritin (µg/mg total protein)

0.87 ± 0.06

0.92 ± 0.05

serum iron (µmol/L)

27.30 ± 5.20

29.60 ± 7.20

serum ferritin (mg/L)

2.10 ± 0.04

2.20 ± 0.15

hemoglobin (g/L)

125.70 ± 8.80

126.00 ± 12.30

serum hepcidin (µg/L)

25.90 ± 11.60

27.50 ± 8.40

Like Hampfl/fl;Myh6.Cre+ mice, Slc40a1 C326Yfl/fl;Myh6.Cre+ mice also had increased mortality relative to their littermate controls (Figure 3C). We then determined whether Slc40a1 C326Yfl/fl;Myh6.Cre+ mice developed the same phenotype of cardiac dysfunction as Hampfl/fl;Myh6.Cre+ mice. Histologically, Slc40a1 C326Yfl/fl;Myh6.Cre+ hearts from six month old mice also showed LV enlargement (Figure 3D), hypertrophied cardiomyocytes (Figure 3E–F), upregulation of hypertrophy markers Myh7 and Nppb (Figure 3G) and a greater degree of apoptosis compared to Slc40a1 C326Yfl/fl controls (Figure 3H–I). When we measured cardiac performance by cine MRI, we found that Slc40a1 C326Yfl/fl;Myh6.Cre+ mice also developed LV dysfunction by 6 months of age, with a reduction in LVEF from 73% to 55% (Figure 3J–L).

The similarity between Hampfl/fl;Myh6.Cre+ and Slc40a1 C326Yfl/fl;Myh6.Cre+ mice in terms of the nature and time course of cardiac dysfunction suggests a common mechanism of cardiac dysfunction involving upregulation of cardiomyocyte FPN. Therefore, we tested whether this upregulation of FPN resulted in increased iron efflux from cardiomyocytes. Iron Fe55 efflux was indeed significantly greater in cardiomyocytes isolated from Hampfl/fl;Myh6.Cre+ and Slc40a1 C326Yfl/fl;Myh6.Cre+ hearts than in cardiomyocytes isolated from their respective controls (Figure 3M). Addition of exogenous mouse HAMP in the efflux medium inhibited the increase in Fe55 efflux from Hampfl/fl;Myh6.Cre+ cardiomyocytes but not from Slc40a1 C326Yfl/fl;Myh6.Cre+ cardiomyocytes, consistent with the HAMP-resistant mutation in Slc40a1 C326Yfl/fl;Myh6.Cre+ cardiomyocytes. We hypothesised that upregulation of cardiac FPN and iron export in Hampfl/fl;Myh6.Cre+ and Slc40a1 C326Yfl/fl;Myh6.Cre+ hearts caused cardiomyocyte iron depletion. To test this hypothesis, we quantified iron levels both in total hearts and in the isolated cardiomyocyte fractions at 3 months and 6 months of age. While iron levels in total hearts were not significantly different between any of the genotypes (Figure 3—figure supplement 2), the iron content of the cardiomyocyte fraction was significantly lower in Hampfl/fl;Myh6.Cre+ and in Slc40a1 C326Yfl/fl;Myh6.Cre+ mice than in their respective controls (Figure 3N). Furthermore, expression of TfR1 mRNA was upregulated (Figure 3O) and Slc40a1 mRNA was downregulated (Figure 3P) in Hampfl/fl;Myh6.Cre+ and Slc40a1 C326Yfl/fl;Myh6.Cre+ hearts relative to their respective controls, consistent with a transcriptional response to intracellular iron deficiency (Rouault and Klausner, 1997; Rouault, 2006; Ward and Kaplan, 1823). Together these results demonstrate that loss of either cardiac hepcidin or hepcidin responsiveness in the heart results in upregulation of cardiomyocyte FPN, and that cardiomyocytes of Hampfl/fl;Myh6.Cre+ and Slc40a1 C326Yfl/fl;Myh6.Cre+ hearts are iron deficient as a result of upregulation of FPN-mediated iron export.

The role of cardiomyocyte iron deficiency and metabolic derangement in cardiac dysfunction

As cardiomyocyte iron deficiency preceded the development of cardiac dysfunction, we hypothesised it is the cause of the cardiac phenotype in Hampfl/fl;Myh6.Cre+ mice. To test this hypothesis, we treated Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl controls with fortnightly intravenous injections of ferric carboxymaltose solution containing 0.5 mg iron from three months of age, and confirmed the effects of this treatment on cardiac and systemic iron indices at 6 months of age (Table 4). At this timepoint, we performed cine MRI and found that the LV enlargement and the reduced LVEF, seen in untreated Hampfl/fl;Myh6.Cre+ mice, were prevented in iron-treated Hampfl/fl;Myh6.Cre+ mice (Figure 4A–C). The transcriptional response to intracellular iron deficiency in untreated Hampfl/fl;Myh6.Cre+ mice (upregulation of TfR1 mRNA and downregulation of Slc40a1 mRNA relative to Hampfl/fl controls), was absent in iron-treated Hampfl/fl;Myh6.Cre+ mice, consistent with correction of cardiomyocyte iron deficiency (Figure 4D–E). Prevention of cardiac dysfunction in Hampfl/fl;Myh6.Cre+ mice by intravenous iron treatment confirms the causal relationship between cardiomyocyte iron deficiency and cardiac dysfunction in this setting.

The role of cardiomyocyte iron deficiency and metabolic derangement in cardiac dysfunction.

(AC) Cine MRI measurements of LV lumen, at end-systole (LVES, *p=0.048), end-diastole (LVED, *p=0.031), and of ejection fraction (LVEF, *p=0.004) in 6-month old untreated (-iron) and I.V iron-treated (+iron) Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. n = 5 per group. (DE) Relative TfR1 (*p=0.001) and Fpn (*p=0.002) expression in hearts of 6-month old untreated (-iron) and I.V iron-treated (+iron) Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. n = 4 per group. (FH) Enzymatic activities of Aconitase I (*p=0.035, 0.041), Complex I (*p=0.004, 0.030) and Complex IV (*p=0.003, 0.026) in untreated (-iron) 3-month and 6-month old and in I.V iron-treated (+iron) 6-month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. n = 4 per group. (I) Representative EM micrographs of hearts from untreated (-iron) 3-month and 6-month old and I.V iron-treated (+iron) 6-month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. Scale bar = 2 µm. (JL) Relative Hk2 (*p=0.010, 0.002), Eno (*p=0.014, 0.021) and Ldha (*p=0.003, 0.001) expression levels in hearts of untreated (-iron) 3-month and 6-month old and in I.V iron-treated (+iron) 6-month old Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. n = 4 per group. NS=not significant. All values are plotted as mean ± SEM.

https://doi.org/10.7554/eLife.19804.019
Table 4

Effect of intravenous iron treatment on iron indices.

Total cardiac and liver elemental iron, serum iron and circulating HAMP in 6-month old untreated and I.V iron-treated Hampfl/fl;Myh6.Cre+ mice and Hampfl/fl littermate controls. Treated mice were injected with 0.5 mg iron fortnighly from the age of 3 months. Tissues and serum were harvested 12 hr after the final injection. n = 5 per group. *p<0.05 relative to untreated Hampfl/fl mice. †p < 0.05 relative to untreated Hampfl/fl;Myh6.Cre+. Values are shown as mean ± SEM.

https://doi.org/10.7554/eLife.19804.021

Hampfl/fl

Hamp fl/fl;Myh6.Cre+

untreated

treated with I.V iron

untreated

treated with I.V iron

cardiac total elemental iron (ng/mg tissue)

82.2 ± 16.9

331.3 ± 21.5*

74.9 ± 7

399.8 ± 68.5†

liver total elemental iron (ng/mg tissue)

100.4 ± 11

2527.6 ± 27.63*

96.3 ± 14

2258.2 ± 239.9†

serum iron (µmol/L)

30.09 ± 6.37

74.48 ± 17.96*

31.5 ± 6.9

80.12 ± 24.9†

serum hepcidin (µg/L)

27.41 ± 6.7

237.3 ± 16.7*

28.9 ± 9.4

209.8 ± 38.8†

Having confirmed a causal relationship between cardiomyocyte iron deficiency and cardiac dysfunction, we aimed to understand the mechanisms linking the two. Iron is a cofactor for several enzymes involved in metabolism (Meyer, 2008; Sono et al., 1996; Solomon et al., 2003), and metabolic derangement is a well-recognized precursor to cardiac dysfunction (Belke et al., 2000; Kakinuma et al., 2000; Ashrafian et al., 2007). Iron deficiency has been reported to reduce the levels and/or activities of key metabolic iron-containing enzymes, in cell lines, in the hearts of mice with impaired cardiac iron uptake (cardiac-specific TfR1 knockouts) and in the hearts of mice fed an iron-deficient diet (Dallman, 1986; Dhur et al., 1989; Xu et al., 2015; Oexle et al., 1999). Based on those studies, we postulated that iron deficiency in the cardiomyocytes of Hampfl/fl;Myh6.Cre+ mice would also result in reduction in the activities of key metabolic iron-containing enzymes. To test this hypothesis, we measured the activities of the iron-sulphur containing enzyme Aconitase I as well as electron transport chain (ETC) complexes in cardiac lysates from Hampfl/fl;Myh6.Cre+ hearts at 3 months and 6 months of age. We found that Aconitase I, Complex I and Complex IV activities were significantly reduced in Hampfl/fl;Myh6.Cre+ hearts compared to Hampfl/fl controls at 3 months and 6 months of age, and that this reduction in activity was prevented in iron-treated 6-month old mice (Figure 4F–H). As ETC activity is essential to mitochondrial function, we examined whether Hampfl/fl;Myh6.Cre+ hearts had signs of mitochondrial failure, and whether such mitochondrial failure was prevented by intravenous iron supplementation. By electron microscopy (EM), dilation of mitochondrial cristae was seen in Hampfl/fl;Myh6.Cre+ hearts as early as 3 months of age and progressed further at 6 months of age. However, this was prevented in iron-treated Hampfl/fl;Myh6.Cre+ mice, which had healthy-looking mitochondria at 6 months of age (Figure 4I). Similar changes in mitochondrial morphology were seen in the hearts of 3 month old Slc40a1 C326Yfl/fl;Myh6.Cre+ mice (data not shown).

Impairment of electron transport is known to drive glycolysis, as an alternative route of ATP production (Schönekess et al., 1997; Kato et al., 2010). Therefore, we examined the expression of a number of genes encoding glycolytic enzymes. We found that expression of genes encoding Hexokinase 2 (Hk2) which catalyses the first step of glycolysis, Enolase (Eno), which catalyses the penultimate step of glycolysis and of Lactate dehydrogenase A (Ldha) which catalyses the ultimate step of glycolysis were all significantly increased at 3 months and 6 months of age in hearts of untreated Hampfl/fl;Myh6.Cre+ mice. This upregulation of glycolytic genes was not seen in iron-treated Hampfl/fl;Myh6.Cre+ mice (Figure 4J–L). These results demonstrate that reduction in the activities of key iron-containing metabolic enzymes, mitochondrial failure and upregulated glycolysis precede the development of cardiac dysfunction in Hampfl/fl;Myh6.Cre+ hearts, and are prevented by intravenous iron treatment.

Discussion

The major finding of this study is that cardiomyocyte hepcidin is required for autonomous cellular iron homeostasis. Loss of hepcidin responsiveness specifically in cardiomyocytes engendered the same effects as loss of cardiac hepcidin, demonstrating that cardiac hepcidin operates in an autocrine fashion by regulating cardiomyocyte FPN. A role in cellular iron homeostasis, and an autocrine mode of action for the HAMP/FPN axis have not been described previously in any other tissue. Indeed, hepcidin and FPN are better known to interact in an endocrine fashion, at the level of the gut, spleen and liver, to regulate systemic iron homeostasis.

A second important finding is that this cardiac HAMP/FPN axis is essential for normal heart function, and that its disruption leads to ultimately fatal cardiac metabolic and contractile dysfunction, even against a background of intact systemic iron homeostasis. Metabolic and contractile dysfunction are preceded by cardiomyocyte iron deficiency and prevented by intravenous iron supplementation, indicating a causal relationship between cardiomyocyte iron deficiency and cardiac dysfunction. This causal relationship has previously been demonstrated in studies using mice lacking cardiac TfR1, in which cardiac iron deficiency also affects the heart against a background of otherwise intact systemic iron homeostasis (Xu et al., 2015). Furthermore, the importance of metabolic derangement seen in our mouse models is also consistent with the findings in mice lacking the cardiacTfR1, and in mice and rats fed iron deficient diets (Dhur et al., 1989; Xu et al., 2015; Tanne et al., 1994; Walter et al., 2002). Correction of metabolic and contractile dysfunction by intravenous iron treatment likely involves not only increased iron availability for uptake into cardiomyocytes, but also the effects of increased circulating HAMP on cardiomyocyte FPN.

Surprisingly, we found that, both in vitro and in vivo, cardiac HAMP protein responded differently from its transcript to changes in iron levels. The current understanding of hepcidin regulation is based on studies of hepatic hepcidin. In that setting, release of the active mature HAMP peptide is dependent on cleavage of the propeptide by Furin (Valore and Ganz, 2008). We found that Furin inhibition increased iron export from Hampfl/fl but not from Hampfl/fl;Myh6.Cre+ cardiomyocytes (Appendix 1—figure 1), demonstrating that cardiomyocytes secrete an active HAMP peptide in a Furin-dependent manner. Furthermore, we found that increased HAMP release from iron-deficient cardiomyocytes depended on Furin, and that cardiac Furin itself is upregulated by iron deficiency both in vitro and in vivo. The latter finding is consistent with the reported regulation of Furin by iron deficiency through Hypoxia-Inducible Factors HIFs (Silvestri et al., 2008). These data suggest that differential regulation of Furin by iron may explain the divergent effects of iron on Hamp transcript and HAMP protein. Comprehensive studies using cardiac-specific knockouts of putative regulators will be required to explore formally the regulation or otherwise, of cardiac hepcidin by pathways known to regulate hepatic hepcidin.

The upregulation of cardiac HAMP in mice fed an iron-deficient diet raises the possibility that it may be involved in protecting the heart in the setting of systemic iron deficiency. This hypothesis is supported by the finding that, when provided an iron-deficient diet, Hampfl/fl;Myh6.Cre+ mice exhibited a greater cardiac hypertrophic response than their Hampfl/fl littermate controls (Appendix 1—figure 2).

Intracellular iron levels are dependent both on cellular homeostatic pathways and on systemic iron availability in plasma. Therefore, the interplay between the cardiac and the systemic HAMP/FPN axes is important in determining cardiomyocyte iron levels. Some insight into this interplay is gained from comparing systemic and cardiac mouse models of disrupted iron homeostasis. It is interesting that ubiquitous Hamp knockout (Lakhal-Littleton et al., 2015) and ubiquitous Slc40a1 C326Y knock-in mice (Appendix 1—figure 3), both models of systemic iron overload, do not develop the cardiac dysfunction seen in cardiomyocyte-specific Hamp knockout and cardiomyocyte-specific Slc40a1 C326Y knock-in mice. This suggests that, while upregulation of cardiomyocyte FPN under conditions of normal iron availability (cardiac-specific models described in this study) is detrimental to cardiac function, it is protective under conditions of increased systemic iron availability (systemic models). Previously, we also showed that deletion of cardiomyocyte FPN resulted in fatal cardiomyocyte iron overload, preventable by dietary iron restriction (Lakhal-Littleton et al., 2015). Together, our studies demonstrate that iron levels within cardiomyocytes are a balance between cellular iron efflux which is regulated by the cardiac HAMP/FPN axis, and systemic iron availability which is regulated by the systemic HAMP/FPN axis (Figure 5).

Interplay between systemic and cardiac iron HAMP/FPN axes.

Cardiomyocyte iron content is determined by both systemic iron availability, which is regulated by liver HAMP, and by the cardiac HAMP/FPN axis, which regulates cardiomyocyte iron efflux. In the wild type heart, cardiac HAMP regulates the levels of cardiac FPN and iron release from cardiomyocytes. In this study, we have demonstrated that loss of cardiac HAMP (cardiac Hamp KO) or loss of cardiac HAMP responsiveness (cardiac Slc40a1 C326Y KI) result in cardiomyocyte iron deficiency due to increased cardiomyocyte FPN and iron release. Previously, we also demonstrated that loss of cardiomyocyte FPN caused cardiomyocyte iron overload. In these two sets of conditions, cardiomyocyte iron deficiency and cardiomyocyte iron overload cause cardiac dysfunction. We have also shown that upregulation of cardiac FPN occurs as a result of loss of either systemic HAMP or systemic HAMP responsiveness, and is protective against the otherwise detrimental effects of systemic iron overload.

https://doi.org/10.7554/eLife.19804.022

Iron overload is detrimental to cardiac health, as demonstrated by iron overload cardiomyopathy in hemochromatosis and thalassemia major patients (Gulati et al., 2014). Our model of cardiac iron homeostasis implies that the cardiac HAMP/FPN axis may have a modifying effect on the severity of iron-overload cardiomyopathy. Thus, it would be interesting to explore whether differences in the levels of cardiac FPN and HAMP, possibly due to different local stimulatory and suppressive signals (e.g local inflammation, local ischemia), explain the reported lack of concordance between the degrees of cardiac iron overload and liver iron overload in a significant proportion of hemochromatosis and thalassemia major patients (Anderson et al., 2001; Noetzli et al., 2008).

Iron deficiency is also detrimental to cardiac health. Indeed, systemic iron deficiency correlates with functional and molecular markers of disease severity in patients with chronic heart failure (CHF) (Klip et al., 2013; Comín-Colet et al., 2013),and also appears to contribute to the risk of death after an episode of acute heart failure (Jankowska et al., 2014). Given the high prevalence of iron deficiency in patients with CHF, ranging between 30–50% (Klip et al., 2013; Nanas et al., 2006; Ezekowitz et al., 2003), the European Society of Cardiology recently recommended the assessment of iron deficiency as a comorbidity in CHF. Furthermore, several clinical trials have now established the benefits of intravenous iron supplementation in CHF patients, with or without anaemia (Ponikowski et al., 2015; Anker et al., 2009; Avni et al., 2012). The mechanisms underlying the anaemia-independent effects of iron deficiency and the benefits of intravenous iron in CHF patients are not fully understood. In light of the direct effect of cardiomyocyte iron deficiency on heart function, demonstrated in this and other studies, it would interesting to explore whether systemic iron deficiency in CHF patients is accompanied by cardiomyocyte iron deficiency, and whether correction of the later underlies the benefits of intravenous iron supplementation in non-anaemic patients.

Another open question is whether disruption of the cardiac HAMP/FPN axis contributes to the pathophysiology of heart disease. It has been shown, in the rat model of myocardial infarction (MI), that Hamp mRNA and HAMP protein are elevated in the ischemic myocardium during the acute phase (Simonis et al., 2010). In humans, circulating HAMP was shown to be elevated in the serum within 4 hr of MI, although the tissue source of this hepcidin was not identified (Suzuki et al., 2009). In addition, decreased cardiac HAMP expression has been reported in a transgenic mouse model of dilated cardiomyopathy, where the phenotype was ameliorated following transgenic overexpression of cardiac hepcidin (Zhang et al., 2012). Thus, further studies are warranted in humans to explore formally the role of the cardiac HAMP/FPN axis in the aetiology of heart disease.

Currently, there is considerable interest in targeting the HAMP/FPN axis for the treatment of iron overload and iron deficiency. Our studies suggest that such strategies may also impinge on cardiac iron homeostasis and function. Other than the heart, both FPN and hepcidin are also found in the brain, kidney and placenta (McCarthy and Kosman, 2014; Kulaksiz, 2005; Evans et al., 2011; Rouault, 2013; Moos and Rosengren Nielsen, 2006; Bastin et al., 2006; Wolff et al., 2011). It would be important to establish whether our findings in the heart extend to these tissues.

Materials and methods

Mice

All animal procedures were compliant with and approved under the UK Home Office Animals (Scientific Procedures) Act 1986. Both males and females were used in experiments, with the respective littermate control being of a matching sex.

The strategy for generating cardiac Hamp knockout mice is outlined in Appendix 1—figure 4. Briefly, a targeting vector was designed to introduce a floxed Hamp allele into C57BL/6N mouse ES cells (JM8F6) with exons 2 and 3, which encode the majority of the peptide, flanked by LoxP sites and a line of floxed mice was generated by blastocyst injection of targeted ES cells, as previous described (Lakhal-Littleton et al., 2015). Further breeding with a C57BL/6 Flp recombinase deleter mouse allowed removal of the Neomycin resistance cassette. Cardiac Hamp knockouts were then generated by by crossing Hampfl/fl animals with mice transgenic for Cre recombinase, under the control of cardiomyocyte-specific Alpha Myosin Heavy Chain (Myh6) promoter (B6.FVB-Tg(Myh6-cre)2182Mds/J).The subsequent breeding strategy was designed to produce cardiac Hamp knockouts and homozygous floxed controls (Hampfl/fl;Myh6.Cre+ and Hampfl/fl respectively) in the same litter.

The strategy for generating Cardiac Slc40a1 C326Y knock-in mice is outlined in Appendix 1—figure 5, and further details are provided in the Appendix.

Cine MRI

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Mice were anaesthetized with 2% isofluorane in O2 and positioned supine in a purpose-built cradle. ECG electrodes were inserted into the forepaws, a respiration loop was taped across the chest and heart and respiration signals were monitored using a custom-built physiological motion gating device. The cradle was lowered into a vertical-bore, 11.7 T MR system with a 40 mm birdcage coil (Rapid Biomedical, Würzburg, Germany) and visualised using a Bruker console running Paravision 2.1.1. A stack of contiguous 1 mm thick true short-axis ECG and respiration-gated cine-FLASH images were acquired. The entire in vivo imaging protocol was performed in approximately 60 min. Image analysis was performed using ImageJ (NIH Image, Bethesda, MD). Left ventricular volumes and ejection fractions were calculated from the stack of cine images as described (Lakhal-Littleton et al., 2015).

Dietary iron content

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Unless otherwise stated, animals were provided with a standard rodent chow diet containing 200 ppm iron. In iron manipulation experiments, mice were given an iron-deficient diet (2–5 ppm iron; Teklad TD.99397; Harlan Laboratories), or an iron-loaded diet (5000 ppm iron; Teklad TD.140464) or a matched control diet (200 ppm iron; Teklad TD.08713) from weaning for six weeks.

Isolation of primary adult mouse cardiomyocytes and in vitro treatment

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Adult primary cardiomyocytes were isolated from eight week old C57BL/6 mice. Hearts were cannulated and mounted on a langendorff apparatus, then perfused using a liberase solution for 10 min. After filtration through a 400 µm gauze, cells were cultured in MEM medium containing Hanks salts, L-glutamine and antibiotics. Within 2 hr of cardiomyocyte culture, supernatants were replaced with fresh medium containing 10% Fetal calf serum, with 0.5 mmol/L ferric citrate (FAC) (F3388, Sigma Aldrich) or 100 µmol/L desferroxamine (D9533, Sigma Aldrich) for 8 hr. The Furin inhibitor decanoyl-Arg-Val-Lys-Arg-chloromethylketone (CMK) (N1505, Bachem) was added at a concentration of 50 µmol/L for the duration of DFO and FAC treatment.

Quantitative PCR

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Total RNA extraction and cDNA synthesis were carried out as previously described (Lakhal-Littleton et al., 2015). Gene expression was measured using Applied Biosystems Taqman gene expression assay probes for Slc40a1, Hamp, TfR1, Myh7, Nppb, Ldha, Hk2, Eno and house-keeping gene β-Actin (Life Technologies, Carlsbad, CA). The CT value for the gene of interest was first normalised by deducting CT value for β-Actin to obtain a delta CT value. Delta CT values of test samples were further normalised to the average of the delta CT values for control samples to obtain delta delta CT values. Relative gene expression levels were then calculated as 2-delta deltaCT.

Immunohistochemistry

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Tissues were prepared as described previously (Lakhal-Littleton et al., 2015) and stained with rabbit polyclonal anti-mouse HAMP antibody (ab30760, Abcam, RRID:AB_2115844) at 1/40 dilution, or rabbit polyclonal anti-mouse FPN antibody (MTP11-A, Alpha Diagnostics, RRID:AB_1619475) at 1/200 dilution. Results of control experiments confirming the specificity of the HAMP antibody are shown in Figure 1—figure supplement 5.

HAMP enzyme-linked immunosorbent assay (ELISA)

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HAMP was measured in mouse sera and in cardiomyocyte supernatants using a HAMP ELISA kit (E91979Mu, USCN) according to the manufacturer’s instructions. Results of control experiments confirming that in vitro treatments did not affect HAMP peptide detection by this ELISA kit are shown in Figure 1—figure supplement 6.

DAB-enhanced perls stain

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Formalin-fixed paraffin-embedded tissue sections were deparaffinised using Xylene, then rehydrated in ethanol. Slides were then stained for 1 hr with 1% potassium ferricyanide in 0.1 mol/L HCl buffer. Endogenous peroxidase activity was quenched, then slides were stained with DAB chromogen substrate and counterstained with haematoxylin. They were visualised using a standard brightfield microscope.

Electron microscopy (EM)

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Hearts were dissected and 0.5–1 mm3 slices were fixed by immersion for 2 hr in 2.5% glutaraldehyde in 0.1 mol/L cacodylate buffer and prepared for electron microscopy by standard methods. Briefly, cells were post-fixed in osmium tetroxide (1% w/v in 0.1 mol/L phosphate buffer), stained with uranyl acetate (2% w/v in distilled water), dehydrated through increasing concentrations of ethanol (70–100%) and acetone and embedded in TAAB resin (TAAB, Aldermaston, UK). Ultrathin sections (50–80 nm) were prepared using a Reichert ultracut S microtome and mounted on 200 mesh nickel grids. Sections were lightly counterstained with lead citrate and uranyl acetate and examined with a Jeol transmission electron microscope (JEM-1010, JEOL, Peabody MA).

Fe55 efflux in primary adult cardiomyocytes

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Adult cardiomyocytes were isolated from mice of the desired genotype at 9 weeks of age as described above. Cardiomyocytes were then cultured in 24-well plates at equal densities for 16 hr before the efflux experiment was performed as described (McKie et al., 2000). Briefly, after washing with three times PBS, cells were incubated for 30 min in 200 µl uptake solution (98 mmol/L NaCl, 2.0 mmol/L KCl, 0.6 mmol/L CaCl2, 1.0 mmol/L MgCl2, 1.0 mmol/L ascorbic acid, 10 mmol/L HEPES [pH 6.0] with Tris base, 50 μmol/L Fe55 [NEN, Boston, MA]), then washed three times with PBS and incubated for 30 min with efflux solution (98 mmol/L NaCl, 2.0 mmol/L KCl, 0.6 mmol/L CaCl2, 1.0 mmol/L MgCl2, 10 mmol/L HEPES [pH 7.4] with Tris base, 300 U/ml bovine ceruloplasmin (cp) (Sigma) and 40 μg/ml human apotransferrin (tf) [Sigma]),in the absence or presence of 0.5 µmol/L mouse HAMP peptide (Peptides International). The efflux medium was then removed, the cells washed three times in ice-cold PBS and disrupted by incubation in 100 µl of 10% SDS solution for 10 min.The efflux solution and cell lysates were then transferred into scintillation vials for Fe55 counting. Where Furin inhibition was carried out, CMK was added to the culture medium at 50 µmol/L 2 hr before the efflux experiment was carried out.

Iron quantitation

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Ferritin concentration in serum and in liver lysates was determined using the ferritin ELISA kit (ICL, Inc. Portland). Serum iron levels were determined using the ABX-Pentra system (Horiba Medical, CA). Determination of total elemental iron in the heart was carried out by inductively coupled plasma mass spectrometry (ICP-MS) as described previously (Lakhal-Littleton et al., 2015). Calibration was achieved using the process of standard additions, where spikes of 0 ng/g, 0,5 ng/g, 1 ng/g, 10 ng/g, 20 ng/g and 100 ng/g iron were added to replicates of a selected sample. An external iron standard (High Purity Standards ICP-MS-68-A solution) was diluted and measured to confirm the validity of the calibration. Rhodium was also spiked onto each blank, standard and sample as an internal standard at a concentration of 1 ng/g. Concentrations from ICP-MS were normalised to starting tissue weight.

Isolation of cardiomyocyte and non-cardiomyocyte fractions for iron quantitation

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Following cardiac perfusion, hearts were dissected into small pieces in ice-cold Hanks buffer, and subject to collagenase P digestion at 37C for 1 hr (11213857001, Roche Diagnostics). Following lysis of red blood cells, cell suspensions were passed through a 70 µm sieve, before being labelled using cardiomyocyte isolation kit (130-100-825, Miltenyi Biotec). Separation of cardiomyocyte and non-cardiomyocyte fractions was carried out using MACS magnetic separation system according to the manufacturer’s instructions. Cardiac fractions were lysed immediately for ICP-MS analysis.

Activity assays for aconitase I and electron transport chain (ETC) complexes

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Approximately 10 mg of frozen, crushed tissues were suspended in 200 µl of ice-cold KME buffer (100 mmol/L KCl, 50 mmol/L Mops, 0.5 mmol/L EGTA, pH 7.4), then homogenized by rupturing with a TissueRuptor (Qiagen, UK) over ice. In a plastic cuvette, the cardiac lysate is mixed with assay buffer and slotted into a spectrophotometer. Details of assay buffers and of reaction procedure for each enzyme are detailed in the Appendix.

Statistics

Values are shown as mean ± standard error of the mean (SEM). Comparison of iron indices, enzyme activities and parameters of cardiac function between groups was performed using Student’s T test. p values < 0.05 were deemed as indicating significant differences between groups. Where significant, exact p values for a figure panel are stated in the corresponding figure legend. No explicit power analysis was performed prior to the experiments to determine sample size, since we had no means to reliably estimate the size and variability of the effects of deleting hepcidin on parameters of cardiac function. For Cine MRI assessment of cardiac function, typically 5–11 animals of each genotype were used, with a matching number of littermate controls. For gene expression, iron quantitation and enzyme activity assays from mouse tissues, typically, 3–6 independent biological replicates and matching littermate controls were analysed. Since significant results were obtained from these set of experiments, no further animals were sacrificed. All 'n' values reported refer to independent biological replicates.

Appendix 1

Generation of Slc40a1 C326Yfl/fl mice

A C57BL/6J mouse genomic BAC clone (RP23-183-P22) encompassing the Slc40a1 gene was used for the generation of the targeting vector. A 12 kb region of the RP23-183-P22 encompassing exons 6–8 and surrounding intronic regions was subcloned by gap-repair into a plasmid containing a diphtheria toxin A chain negative selection cassette. The required C326Y point mutation was introduced into this subclone by site directed mutagenesis. In parallel, a loxP flanked neomycin/kanamycin selection cassette (from PL452 Liu et al., 2003) was inserted into the BAC within Slc40a1 intron six by recombineering and an approximately 5 kb ApaI/PmeI fragment encompassing this insertion together with exons 6 and 7 and flanking introns was subcloned into pBluescript. This plasmid was treated with Cre recombinase (NEB) in vitro to remove the selection cassette, resulting in a plasmid containing a single loxP and an additional SpeI site within Slc40a1 intron 6. This loxP interrupted sequence was shuttled into the genomic subclone by exchanging an internal NcoI fragment. A portion of Slc40a1 cDNA corresponding to the coding region encoded on exons 7 and 8, including the 3’ untranslated region was amplified by RT-PCR from cDNA prepared from embryonic mouse heart and inserted into a piece of synthetic DNA (Gene Art) corresponding to intronic and intergenic sequence lying immediately upstream of exon seven and downstream of exon 8, respectively. This minigene assembly was cloned upstream of an FRT-PGK-Neomycin-FRT-loxP selection cassette (from P451 Liu et al., 2003) and the two pieces together were inserted into the modified genomic subclone above via a unique SacII site, completing the final targeting vector. The completed targeting vector was linearised with AatII and electroporated into C57BL/6N JM8F6 embryonic stem cells. Following selection in 210 μg/ml G418, recombinant clones were screened by PCR to detect homologous recombination over the 3’ arm. A forward primer (5’- CAATAGCAGGCATGCTGGGGATG-3’) binding within the polyadenylation of the selection cassette was used together with a reverse primer (5’- GGCATCTGCTGTCTGTGAAA- 3’) binding downstream of the 3’ homology arm to amplify a 6.6 kb fragment from correctly recombined clones. Positive clones were examined for correct recombination at the 5’ end by long range PCR using a forward primer (5’- TGTACCATGGATGGGTCCTT-3’) binding upstream of the 5’ homology arm and a reverse primer (5’- TACCGGTGGATGTGGAATGTG- 3’) binding within the PGK promoter driving the neomycin selection. Correctly targeted clones yielded a 9.7 kb amplicon. Incorporation of the 5’ loxP site, was verified by digestion of this 5’ amplicon with SpeI which yielded a 7 kb and 2.7 kb cleavage products in correctly targeted clones. Both 5’ and 3’ screening amplicons were analysed by Sanger sequencing to confirm the junction sequence over the extremities of the homology arms using sequencing primers 5’-AAAGCCCAGGGGTATCTCTC-3’ and 5’- CTCAGCTTGGCTATGTGGTG-3’ for the 5’ and 3’ amplicons respectively. The presence of the 5’ loxP site within the 5’ amplicon was confirmed by Sanger sequencing using sequencing primer 5’- CATGAAGCAGTGGGCATAGA-3’ and the presence of the C326Y mutation was confirmed within the 3’ amplicon using sequencing primer 5’-CACACACACACATATATACATGCAA-3’. Southern blotting using a probe against neomycin was used to confirm that only a single integration event had occurred. Correctly recombined ES cells were injected into albino C57BL/6J blastocysts and the resulting chimeras were mated with albino C57BL/6J females. Successful germline transmission yielded black pups and F1 mice harboring the conditional C326Y knock-in allele were identified using the above screening PCR. F1 heterozygous male mice were bred with C57BL/6J Flp recombinase deleter mice (Tg(ACTB-Flpe)9205Dym (Jax stock 005703)) and offspring were screened for the deletion of the selection cassette using a forward primer (5’-GATATCATCATCGCCCTTTGG- 3’) binding within the 3’ untranslated region of the exon 7–8 minigene and a reverse primer (5’- TTGCATGTATATATGTGTGTGTGTG-3’) binding immediately downstream of the cassette. A 1.4 kb amplicon was obtained from the Flp deleted conditional C326Y knock-in allele. Heterozygous mice without the selection cassette were then backcrossed with C57BL/6J to remove the Flp transgene prior to onward breeding.

Aconitase and ETC activities

Total aconitase I activity assay

Total aconitase I activity was measured as the rate of isomerization of cis-aconitic to isocitrate at 240 nm and 37°C using a spectrophotometer. The assay buffer consisted of 50 mmol/L Tris HCl pH 7.5. The reaction was carried out in a quartz cuvette containing 1 ml assay buffer supplemented with 10 µl 20 nmol/L cis-aconitic acid (Sigma) and 25 µl of protein lysate and read for 2 min at 240 nm against a black containing ddH2O. The activity of aconitase was determined by dividing the gradient of the absorbance over the extinction coefficient (3600) and expressed in nmol/min/mg tissue.

Complex I activity assay

Mitochondrial complex I activity was measured as the rate of NADH oxidation at 340 nm and 30°C using a spectrophotometer. The assay buffer contained the following: 25 Mmol/L potassium phosphate pH 7.2, 5 mmol/L MgCl2 (Sigma), 0.13 mmol/L NADH (MP Biomedicals, LLP), 65 µM coenzyme Q1 (Sigma), 250 mg fatty acid free BSA (Sigma), 200 µg antimycin A (Santa Cruz). The reaction was carried out in 1 ml assay buffer supplemented with 20 µl of protein lysate and read for 1 min at 340 nm against a blank containing ddH2O. As a negative control, 20 µl of 10 mmol/Lrotenone (Sigma) was added to a new reaction and the inhibited rate measured for 1 min. The activity of complex I was determined by dividing the gradient of the absorbance over the extinction coefficient (6810) and expressed in nmol/min/mg tissue.

Complex II activity assay

Mitochondrial complex II activity was measured as the rate of succinate-dependent reduction of dichlorophenolindophenol (DCIP) at 600 nm and 30°C using a spectrophotometer. The assay buffer contained the following: 25 mol/L potassium phosphate pH 7.2, 5 mmol/L MgCl2, and 2 mmol/L sodium succinate dibasic hexahydrate (Sigma). The reaction was carried out in 985 µl of assay buffer supplemented by 10 µl of buffer containing 2 µg antimycin, 2 µg rotenone, 50 µmol/L DCIP (Sigma) followed by addition of 65 µmol/L CoQ1 and 20 µl protein lysate and read for 2 min at 600 nm against a blank containing ddH2O. As a negative control, 20 µl of 10 mmol/L sodium malonate (Sigma) was added to a new reaction and the inhibited rate measured for 2 min. The activity of complex II was determined by dividing the gradient of the absorbance over the extinction coefficient (21000) and expressed in nmol/min/mg tissue.

Complex III activity assay

Mitochondrial complex III activity was measured as the rate of reduction of cytochrome c3+ at 550 nm at 30°C using reduced ubiquinol as an electron acceptor using a spectrophotometer. To prepare ubiquinol, 10 mg of decylubiquinone (Sigma) was dissolved in 312.5 µl absolute ethanol to give a final concentration of 100 mmol/L. An aliquot of 100 µl decylubiquinone working solution was further diluted in 900 µl ethanol and acidified to pH 2 with a 6 mol/L HCl solution. The ubiquinone was then reduced with a pinch of sodium borohydrate (Sigma) and 1 ml of ddH2O was added to stop the reaction. Excess sodium borohydrate was allowed to settle and the sample centrifuged briefly to separate the sodium borohydrate precipitate from the reduced ubiquinol. Using a pH indicator paper, the pH of the ubiquinol was verified to be at pH two before using in the activity assay. The reaction was carried in 1 ml of assay buffer (50 mM potassium phosphate pH 7.2, 3 mmol/L sodium azide (Sigma), 1.5 µmol/L rotenone, and 50 µmol/L cytochrome c from bovine heart [Sigma]), supplemented with 5 µl reduced ubiquinol and 20 µl protein lysate and read for 1 min at 550 nm against a blank containing ddH2O. As a negative control, 10 µl of 20 mmol/L antimycin A was added to a new reaction and the inhibited rate measured for 1 min. As further negative control, a reaction containing protein only or reduced ubiquinol only was also measured for 1 min. The activity of complex III was determined by dividing the gradient of the absorbance over the extinction coefficient (19100) and expressed in nmol/min/mg tissue.

Complex IV activity assay

Mitochondrial complex IV activity was measured as the rate of oxidation of cytochrome c2+ at 550 nm at 30°C using a spectrophotometer. To prepare the reduced cytochrome c2+, a dialysis tubing (Viking 7000/1) of appropriate length was hydrated for 30 min in 1 L ddH2O supplemented with 20 g sodium carbonate and 0.372 g EDTA at 80°C. A solution of 100 mg of cytochrome c from bovine heart and 10 mg sodium ascorbate dissolved in 10 ml potassium phosphate (0.1 mol/L, pH 7.0) was added to the tubing and dialysed against 1 L of 0.1 mol/L potassium phosphate buffer for 24 hr at 4°C. The phosphate buffer was exchanged three times, every 8 hr. The redox state of the synthesized reduced cytochrome c was verified by measuring the absorbance spectra between wavelengths of 500 and 600 nm, in the presence or absence of 100 µmol/L potassium ferricyanide (Sigma) and compared against the oxidized cytochrome c. The reaction was carried out in 1 ml assay buffer (10 mmol/L potassium phosphate pH 7.0 and 0.50 µmol/L reduced cytochrome c) supplemented with 5 µl protein lysate and read for 3 min at 550 nm against a blank of assay buffer supplemented by 100 µmol/L potassium ferricyanide. As a negative control, 10 µl of 10 mmol/L sodium azide was added to a new reaction and the inhibited rate measure for 3 min. The first-order rate constant (k) was calculated as previously described Brooks et al. (2000). Briefly, the natural logarithm was taken for the absorbances at four time points t = 0, 60, 120, 180 s and the difference for each pair of time points determined (t(60)-t(0), t(120)-t(60), t(180)-t(120)). The average of these differences were taken to be k and the activity expressed in nmol/min/mg tissue.

Appendix 1—figure 1
Effect of Furin inhibitor on iron export in cardiomyocytes.

Fe55 efflux measured in primary adult cardiomyocytes from Hamp fl/fl and Hamp fl/fl;Myh6.Cre+ mice following culture in control medium (-CMK) or medium containing Furin inhibitor (+CMK) for 2 hr. n = 3. values are plotted as mean ± SEM. *p=0.027. †p = 0.024.

https://doi.org/10.7554/eLife.19804.023
Appendix 1—figure 2
Effect of genotype on hypertrophic response to iron-deficient diet.

Heart/body weight ratio in Hampfl/fl;Myh6.Cre+ mice and littermate Hampfl/fl controls provided a control diet (Fe 200 ppm) or an iron-deficient diet (Fe 2–5 ppm) from weaning for six weeks. n = 5 per group, NS=not significant. *p=0.013, †p = 0.04. Values are shown as mean ± SEM.

https://doi.org/10.7554/eLife.19804.024
Appendix 1—figure 3
Normal cardiac function in ubiquitous Slc40a1 C326Y KI mice.

(A–C) Cine MRI measurements of LV lumen, at end-systole (LVES), end-diastole (LVED), and of ejection fraction (LVEF) in Slc40a1 C326YKI/+ mice and +/+ littermate controls at three months (n = 7 per group), six months (n = 7 per group) and nine months (n = 7 per group) of age. Values are plotted as mean ± SEM.

https://doi.org/10.7554/eLife.19804.025
Appendix 1—figure 4
Strategy for generation of Hampfl/fl;Myh6.Cre+ mice.

A targeting vector was designed to introduce a floxed Hamp allele into mouse ES cells, with exons 2 and 3, which encode the majority of the peptide, flanked by LoxP sites. Further breeding with a C57BL/6 Flp recombinase deleter mouse allowed removal of the Neo cassette. Cardiac Hamp knockouts were then generated by crossing homozygous Hampfl/fl animals with mice transgenic for Myh6-Cre recombinase, which is under the control of cardiomyocyte-specific Myosin Alpha Heavy chain six promoter.

https://doi.org/10.7554/eLife.19804.026
Appendix 1—figure 5
Strategy for generation of Slc40a1 C326Yfl/fl;Myh6.Cre+ mice.

A targeting vector was designed to introduce a floxed Slc40a1 C326Y allele into mouse ES cells, containing mutant exon seven and wild type and to delete simultaneously endogenous exons 7 and 8. Further breeding with a C57BL/6 Flp recombinase deleter mouse allowed removal of the Neo cassette. Cardiac Slc40a1 C326Y knock-ins were then generated by crossing homozygous Slc40a1 C326Yfl/fl animals with mice transgenic for Myh6-Cre recombinase, which is under the control of cardiomyocyte-specific Myosin Alpha Heavy chain six promoter.

https://doi.org/10.7554/eLife.19804.027
Appendix 1—figure 6
Additional images for HAMP and FPN staining.

(A) Lower magnifications images (X10) for Figure 1B. (B) Lower magnification images (X10) for Figure 1F. (C) Higher magnification images (X40) for Figure 3A and C.

https://doi.org/10.7554/eLife.19804.028

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Decision letter

  1. Richard P Harvey
    Reviewing Editor; The Victor Chang Cardiac Research Institute, Australia

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "An essential cell-autonomous role for hepcidin in cardiac iron homeostasis" for consideration by eLife. Your article has been favorably evaluated by Naama Barkai (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

The reviewers found the manuscript to have merit and potentially to make an important contribution to understanding iron homeostasis at the tissue level. For the first time, the authors tested the conjecture that tissue hepcidin production may contribute to local tissue-level iron regulation.

However, a number of important issues were raised by the reviewers including the need to be stricter in the methodology used and its description. In particular, the reviewers were seeking further evidence that the data on hepcidin expression and regulation were robust and that bioactive hepcidin was being detected, with hepcidin KOs as controls. Reviewers were reluctant to accept immunohistochemical staining as quantification of bioactive secreted hepcidin. Furthermore, iron efflux experiments need to be described in more detail. Tissue heterogeneity including the possible presence of macrophages may be an issue to resolve. Any further evidence that can be provided on the regulation of hepcidin would strengthen the paper.

I have summarised the major issues below:

1) Subsection “Hamp expression and regulation in the heart”, first paragraph and Figure 1: The semiquantitation of hepcidin by immunohistochemical methods has serious liabilities because hepcidin is a secreted protein and because antibodies can detect functional and nonfunctional hepcidin peptide, including precursor and degraded forms. The conversion of propeptide to active peptide is a step that could be regulated by iron and this may not be detected by the method used. The Methods section indicates that it was a rabbit polyclonal anti-mouse antibody from Abcam, but it does not say which antibody from Abcam was used. Several listed on Abcam's website were prepared against human peptides but have anti-mouse reactivity; one was prepared against mouse antigen and recognizes both Hamp1 and Hamp2 in mice. Are the authors certain that the antibody is recognizing the hepcidin-25 peptide encoded by Hamp1, and not something else? If they are detecting either pro-hepcidin or Hamp2 the conclusions of the paper might be very different. Also, because hepcidin functions as an extracellular peptide, it is difficult to draw conclusions from intracellular antibody staining. Specifically: a) It is not clear which hepcidin forms are stained and where they are located. Does the antibody detect intracellular peptide which is mostly propeptide? Fluorescent confocal microscopy could help with localization. b) No evidence is provided that cardiac myocytes can generate the functional 25 amino acid hepcidin. The processing of peptide hormones to bioactive forms requires prohormone convertases which are expressed in a tissue-specific manner.

2) Subsection “Hamp expression and regulation in the heart”, first paragraph and Figure 1D: The detection of hepcidin secreted by cardiomyocytes in culture needs to be validated better because the concentrations are very low, and because the post-mRNA effect of DFO and FAC on hepcidin secretion is unprecedented. This experiment is crucial for understanding the function of local control by hepcidin-ferroportin. It is therefore necessary to be as careful as possible here. Controls should include hepcidin KO cardiomyocytes. At these low peptide concentrations, adsorption of peptide to plastic surfaces causes major losses and these could be affected by DFO and FAC treatment. Neither is it clear which forms of hepcidin are released into the media. Finally, primary hepatocytes within less than 24h in culture lose their ability to physiologically regulate hepcidin in response to iron, and this can result in delayed paradoxical suppression of hepcidin mRNA by FAC. If this happens with cardiomyocytes it could lead to artefacts. The regulation of hepcidin mRNA by FAC and DFO should therefore be checked as a function of time in culture.

3) To confirm the equivalence of Hamp KO and ferroportin C326Y the survival curves need to be compared. This is a hard endpoint that has produced a surprise previously (e.g. the marked difference in survival between global hepcidin KO and global ferroportin C326S). If data exists on the survival of Ferroportin C326Y transgenic mice, it should be presented.

4) It would be relatively straightforward for the authors to examine the regulation of cardiac Hamp using hemojuvelin floxed (Hfe2 fl/fl) mice (Chen W et al., Blood 2011) or other approaches, as regulation of hepatic Hamp expression is well understood. Hfe2 is expressed at high levels in the heart and might be relevant there as it is in the liver. The paper would be much stronger if there was some indication of the mechanism of hepcidin regulation in cardiomyocytes.

5) The authors refer to other animal models of cardiac iron deficiency, but they don't really distinguish between those caused by dietary iron deprivation, which would have the confounding existence of iron deficiency anemia, and a heart-specific Tfrc knockout model in which iron deficiency is restricted to the heart (Xu W et al., Cell Reports 2015, ref 31). The heart-specific Tfrc knockout was shown to have many characteristics that are similar to those described in this paper. One important difference, however, was in the levels of electron transport chain complexes found in untreated and iron treated control and cardiac Hamp-/- mice (Figure 4—figure supplement 2). Xu et al. reported decreases in the bands representing iron-containing ETC complexes using the same OXPHOS blotting approach, as would be expected in iron deficient cells. Those representative proteins are labile when the complexes do not form properly. How do the authors explain their unexpected results, particularly considering the decreased complex activity shown in Figure 4?

6) Several experiments were carried out using isolated adult mouse cardiomyocytes. How did the authors insure that the cell populations were not contaminated with other cell types, particularly macrophages, which might confound the results?

7) The iron efflux experiments shown in Figure 3L appear to be novel because there is no reference provided. Is this correct? For that reason it would be very helpful to know more about them, to assess their validity. Apparently, cells were loaded with non-transferrin bound 55Fe by a 30 minute incubation. How much iron was taken up under these conditions? Why was non-transferrin bound iron used instead of Fe-Tf, particularly considering the results described in Xu et al., 2015 indicating that the transferrin cycle is very important for cardiomyocyte iron uptake? What measures were undertaken to ensure that the PBS washes effectively removed non-assimilated radiolabeled iron? How many times were the experiments repeated, and how variable were the results? Why do the authors think the FpnC326Y mutant had higher iron efflux compared to Hamp-/- cardiomyocytes in the absence of Hamp?

8) Figure 1, panel A – this appears to be mRNA expression but this is not stated. If it is, how was it normalized? In panel B, as the authors pointed out, it is quite surprising that there appears to be most anti-Hamp staining under iron deficient conditions. If this is truly physiologically important staining (see point 2 above) then it becomes even more important to understand how Hamp expression is regulated (see point 3 above). Hamp is typically measured extracellularly, rather than intracellularly because it is a secreted protein; a defect in pre-protein processing might help explain the discrepancy between panels B and C in this figure. Panel D is particularly important – how many times was this experiment replicated? How reliable is the assay kit (the methods indicate that it is from "uscnk" but I don't know what that is – please specify), and is it specific for Hamp1 rather than Hamp2?

9) The heart sections in Figure 2B are suboptimal. It is particularly difficult to discern anatomy in the fl/fl section. In Figure 2D, the control cardiomyocytes were much smaller than those reported in Figure 1G of Xu et al., 2015, and the difference in cardiomyocyte surface area between controls and mutants was much larger. Do the authors have any idea what this difference might be due to?

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your article "An essential cell-autonomous role for hepcidin in cardiac iron homeostasis" for consideration by eLife. Your article has been favorably evaluated by Naama Barkai (Senior Editor) and a Reviewing Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

The reviewers generally did not think that the authors use of tissue immunostaining to quantitate a secreted protein is bullet-proof but accept that the authors have enough additional overlapping evidence to give more confidence that the main conclusions are right. They have put together a strong case for hepcidin and ferroportin interacting in an autocrine/paracrine manner in the heart. The genetic argument from the equivalent pathology and similar survival of cardiomyocyte FpnC326Y (cardiomyocyte hepcidin resistance) and cardiomyocyte hepcidin KO is particularly appealing. The autocrine/paracrine function of hepcidin and ferroportin is an old conjecture but this is the first evidence for it.

However, the following is required for acceptance:

1) In the rebuttal there is a statement that the reviewers did not agree with. The cardiac-specific Tfr1 KO mice are not embryonic lethal – they live into the second postnatal week. The authors should be aware of this and modify the text if needed.

2) One reviewer did not believe that the authors had fully explained the outstanding issues with their OXPHOS blot – they thought it was poor quality with much more animal to animal variability than there should be. If possible, the authors should repeat this blot in order to show more consistency between replicates. Alternatively, the authors should delete this western.

https://doi.org/10.7554/eLife.19804.029

Author response

[…]

1) Subsection “Hamp expression and regulation in the heart”, first paragraph and Figure 1: The semiquantitation of hepcidin by immunohistochemical methods has serious liabilities because hepcidin is a secreted protein and because antibodies can detect functional and nonfunctional hepcidin peptide, including precursor and degraded forms. The conversion of propeptide to active peptide is a step that could be regulated by iron and this may not be detected by the method used. The Methods section indicates that it was a rabbit polyclonal anti-mouse antibody from Abcam, but it does not say which antibody from Abcam was used. Several listed on Abcam's website were prepared against human peptides but have anti-mouse reactivity; one was prepared against mouse antigen and recognizes both Hamp1 and Hamp2 in mice. Are the authors certain that the antibody is recognizing the hepcidin-25 peptide encoded by Hamp1, and not something else? If they are detecting either pro-hepcidin or Hamp2 the conclusions of the paper might be very different. Also, because hepcidin functions as an extracellular peptide, it is difficult to draw conclusions from intracellular antibody staining. Specifically: a) It is not clear which hepcidin forms are stained and where they are located. Does the antibody detect intracellular peptide which is mostly propeptide? Fluorescent confocal microscopy could help with localization. b) No evidence is provided that cardiac myocytes can generate the functional 25 amino acid hepcidin. The processing of peptide hormones to bioactive forms requires prohormone convertases which are expressed in a tissue-specific manner.

Hepcidin is synthetized as an 84 amino acid prepropeptide, containing an N-terminal ER targeting signal sequence (aa 1-24), a proregion (pro) with a consensus Furin cleavage site (aa 25-59) and a C-terminal bioactive iron-regulatory mature peptide (aa 60-84). The antibody used for immunohistochemical detection of hepcidin in the heart is ab30760 from Abcam, which was raised against the human mature peptide (aa60-84). We now include this information in the Methods section of the revised manuscript.

The reviewers’ concerns over the specificity of this antibody are ones that we also had, and consequently we did carry out appropriate control experiments. We now include the results of those control experiments in the revised manuscript. First, we showed that staining with this antibody is absent in the heart of cardiac-specific hamp-1 knockouts (Figure 1F), despite Hamp-2 expression being unaltered in these hearts compared to controls (Figure 1—figure supplement 5). These data demonstrate that this antibody detects HAMP1 and not HAMP2. Second, we showed that staining in the heart of wild-type animals is completely abrogated by the use of the mature peptide (hepcidin-25, Abcam ab 31875) as a blocking peptide (Figure 1—figure supplement 5). In hepatocytes, it has been shown that the propeptide can also be secreted, and that this propeptide is detectable by antibodies raised against the mature peptide (Rouault and Klausner, 1997). Thus, in theory our antibody detects not only the mature secreted peptide but also the propeptide, and even demonstration of extracellular staining by immunohistochemistry would not constitute formal evidence for the presence of the active mature peptide. We do, however, provide direct evidence for the secretion of an active hepcidin by cardiomyocytes. First, in our in vitro experiments, where we detect hepcidin by ELISA in supernatants of primary cardiomyocytes from control animals (original data). We reproduced this finding in a second independent experiment (new Figure 1D), and further show that hepcidin is greatly reduced or undetectable in supernatants of cardiomyocytes from cardiac Hamp-1 knockout animals. We now include these data as Figure 1—figure supplement 4. Second, we also provide evidence that the secreted peptide is active, and this is discussed below.

The reviewer is right to point out that the release of the bioactive hepcidin mature peptide requires prohormone convertases, which may or may not be present in the heart. Furin is the prohormone convertase required for release of hepatic bioactive mature hepcidin peptide. In hepatocytes, it has been shown that Furin inhibition leads to decreased release of the active mature peptide and increased release of the larger inactive propetide, and that both of these protein species are detected by assays targeting the mature peptide (Rouault and Klausner, 1997). Furin is also highly expressed in the heart (Beaubien et al., 1995), raising the intriguing possibility that it may be involved in the production of the bioactive hepcidin mature peptide from cardiomyocytes. To test this hypothesis, we performed additional experiments using the Furin inhibitor Decanoyl-Arg-Val-Lys-Arg-chloromethylketone trifluoroacetate salt(CMK). These experiments showed that while Furin inhibition under iron replete conditions did not decrease secreted HAMP levels (consistent with the release of the propeptide as reported in hepatocytes) (Valore and Ganz, 2008), it nonetheless increased iron export from primary cardiomyocytes. Furthermore, the effect of Furin inhibitor on iron export was only seen in Hamp fl/fl and not in Hamp fl/fl Myh6.Cre+ cardiomyocytes. We include these data in Appendix 1—Figure 1. Together, these results provide evidence that cardiomyocytes release an active form of hepcidin in a Furin-dependent manner.

Furthermore, we found that increased HAMP release from DFO-treated cardiomyocytes depended on Furin. These data are added to Figure 1D. We also found that Furin itself was upregulated by iron deficiency in vitro and in vivo. We include these data as Figure 1—figure supplement 2. These findings are consistent with the previously reported regulation of Furin by iron deficiency through Hypoxia-inducible factors HIFs (Silvestri, Pagani and Camaschella, 2008). They suggest that differential regulation of Furin by iron may explain the divergent effects of iron on Hamp transcript and HAMP protein.

2) Subsection “Hamp expression and regulation in the heart”, first paragraph and Figure 1D: The detection of hepcidin secreted by cardiomyocytes in culture needs to be validated better because the concentrations are very low, and because the post-mRNA effect of DFO and FAC on hepcidin secretion is unprecedented. This experiment is crucial for understanding the function of local control by hepcidin-ferroportin. It is therefore necessary to be as careful as possible here. Controls should include hepcidin KO cardiomyocytes. At these low peptide concentrations, adsorption of peptide to plastic surfaces causes major losses and these could be affected by DFO and FAC treatment. Neither is it clear which forms of hepcidin are released into the media. Finally, primary hepatocytes within less than 24h in culture lose their ability to physiologically regulate hepcidin in response to iron, and this can result in delayed paradoxical suppression of hepcidin mRNA by FAC. If this happens with cardiomyocytes it could lead to artefacts. The regulation of hepcidin mRNA by FAC and DFO should therefore be checked as a function of time in culture.

As suggested by the reviewer, we have performed additional experiments with primary cardiomyocytes from cardiac hamp1-ko cells, and found that levels of HAMP proteins were greatly reduced or undetectable in their supernatants. We include this in Figure 1—figure supplement 4. Furthermore, the concentrations detected in the supernatants are well within the detection range (4.94-400pg/mL) of the ELISA kit used (Uscn Life Science Inc. #E91979Mu). The reviewer was also concerned that in vitro treatments (DFO and FAC) may affected detection of the hepcidin peptide by this ELISA assay. To guard against the possibility of differential absorption resulting from the presence of DFO or FAC, we compared the standard curves obtained by dilution of the peptide standard in a) standard diluent provided in the kit, b) unconditioned cardiomyocyte growth medium, c) unconditioned cardiomyocyte growth medium with 100uM DFO, and d) unconditioned cardiomyocyte growth medium with 500uM FAC. We found no significant differences between the 4 standard curves. We include these data as Figure 1—figure supplement 6. This demonstrates that the presence of DFO or FAC does not affect the detection of the hepcidin peptide by this ELISA kit.

The reviewer was concerned that physiological control of hepcidin was lost in cardiomyocytes due to the length of culture. First, both in the original and in the additional experiments performed herein, cardiomyocytes were used within 2 hours of harvest. Additionally, as suggested by the reviewer, we carried out a timecourse of treatment with FAC and DFO, and now present these data in the manuscript (new Figure 1C for Hamp transcript and new Figure 1D for HAMP in supernatants). Overall, the direction of change in response to DFO and FAC, both for hepcidin transcript and protein are maintained over the 24 hour period and are consistent with the single-timepoint data presented in the original manuscript prior to revision.

3) To confirm the equivalence of Hamp KO and ferroportin C326Y the survival curves need to be compared. This is a hard endpoint that has produced a surprise previously (e.g. the marked difference in survival between global hepcidin KO and global ferroportin C326S). If data exists on the survival of Ferroportin C326Y transgenic mice, it should be presented.

We now provide those data as new Figure 3C. As for Hamp fl/fl;Myh6.Cre+ mice, Slc40a1 fl/fl;Myh6.Cre+ animals also have greater mortality compared to their littermate controls with mean survival of 32 weeks.

4) It would be relatively straightforward for the authors to examine the regulation of cardiac Hamp using hemojuvelin floxed (Hfe2 fl/fl) mice (Chen W et al., Blood 2011) or other approaches, as regulation of hepatic Hamp expression is well understood. Hfe2 is expressed at high levels in the heart and might be relevant there as it is in the liver. The paper would be much stronger if there was some indication of the mechanism of hepcidin regulation in cardiomyocytes.

HJV is one of many putative regulators of cardiac hepcidin. As it happens, muscle-specific HJV KO mice have increased rather than decreased cardiac hepcidin expression (Kostas Pantopoulos, personal communication). Thus, it appears that both with respect to HJV and to iron, cardiac hepcidin is regulated differently from liver hepcidin. Substantive additional studies are required to explore the role of known regulators of hepatic hepcidin in the regulation of cardiac hepcidin. We consider such studies beyond the scope of this revision as the work would constitute an independent manuscript.

5) The authors refer to other animal models of cardiac iron deficiency, but they don't really distinguish between those caused by dietary iron deprivation, which would have the confounding existence of iron deficiency anemia, and a heart-specific Tfrc knockout model in which iron deficiency is restricted to the heart (Xu W et al., Cell Reports 2015, ref 31). The heart-specific Tfrc knockout was shown to have many characteristics that are similar to those described in this paper. One important difference, however, was in the levels of electron transport chain complexes found in untreated and iron treated control and cardiac Hamp-/- mice (Figure 4—figure supplement 2). Xu et al. reported decreases in the bands representing iron-containing ETC complexes using the same OXPHOS blotting approach, as would be expected in iron deficient cells. Those representative proteins are labile when the complexes do not form properly. How do the authors explain their unexpected results, particularly considering the decreased complex activity shown in Figure 4?

With regards to the description of animal models of cardiac iron deficiency, we have now amended the text to distinguish between the various models, as suggested by the reviewer. With regards to the levels of ETC complexes detected by OXPHOS, we too were initially surprised that we did not detect a reduction as was seen in cardiac TfR knockout mice (Xu et al., 2015).However, this becomes less surprising when you consider the relative severity of cardiac iron deficiency, which is reflected in the fact that cardiac-specific TfR KO are embryonic lethal whereas cardiac-specific hepcidin knockouts have a mean survival of 28 weeks. In this context, it is important to consider the mechanisms through which iron deficiency impinges on the ETC. These mechanisms include not only limiting iron availability for complex assembly and for enzymatic activity of assembled complexes, but also reducing the levels of mitochondrially-encoded proteins. Indeed, the degree of iron deficiency (and iron excess) have been shown to correlate with damage to mitochondrial DNA (Walter et al., 2002), and many of the proteins in ETC complexes are encoded by mitochondrial genes. Thus, depending on the degree of iron deficiency in the cardiomyocytes, different mechanisms may come into play, meaning that complex assembly or activity could still be disrupted when levels of individual proteins in the complex are still maintained.

6) Several experiments were carried out using isolated adult mouse cardiomyocytes. How did the authors insure that the cell populations were not contaminated with other cell types, particularly macrophages, which might confound the results?

The reviewer is right to point out that macrophages are known to be present in the cardiac tissue. We cannot exclude the presence of macrophages in our cardiomyocyte culture. However, specifically with respect to hepcidin production in the supernatants, we can be confident that cardiomyocytes are the source of the hepcidin, because very little hepcidin was detected in supernatants of cardiac-specific hepcidin knockouts (where cardiomyocytes but not macrophages have deletion of the hepcidin gene). We include these data in Figure 1—figure supplement 4.

7) The iron efflux experiments shown in Figure 3L appear to be novel because there is no reference provided. Is this correct? For that reason it would be very helpful to know more about them, to assess their validity. Apparently, cells were loaded with non-transferrin bound 55Fe by a 30 minute incubation. How much iron was taken up under these conditions? Why was non-transferrin bound iron used instead of Fe-Tf, particularly considering the results described in Xu et al., 2015 indicating that the transferrin cycle is very important for cardiomyocyte iron uptake? What measures were undertaken to ensure that the PBS washes effectively removed non-assimilated radiolabeled iron? How many times were the experiments repeated, and how variable were the results? Why do the authors think the FpnC326Y mutant had higher iron efflux compared to Hamp-/- cardiomyocytes in the absence of Hamp?

The method for measuring iron efflux used in our experiments is not novel, but adapted from the method used by Mckie et al. 2000, which was the first publication to describe the function of FPN in iron efflux. We now include this reference in the text. This method does not enable measurement of absolute amounts of iron flux, but rather measurement of iron efflux as a percentage of the total iron taken up during incubation with the uptake solution; percentage efflux= 100 x Fe55 in efflux solution /(Fe55 in efflux+ Fe55 remaining in cells). The readouts are obtained using a scinitillation counter and are expressed as counts per minute. Count rate does not universally equate to dose rate, and there is no simple universal conversion factor. Any conversions are instrument-specific.

The reviewer also questions the use of non-transferrin bound Fe55 (as in Mckie et al., 2000) instead of transferrin-bound Fe55 for loading cells with iron. The rationale for this is to circumvent any confounding effects of differential iron uptake between cells of different genotypes due to differing levels of TfR1 expression. Indeed, we know that TfR1 expression is higher in cardiac-hepcidin knockout cells and in Slc40a1 C326Y knock-in cells than in their respective controls (Figure 3O). While TfR1 is the dominant route of iron uptake into cardiomyocytes, it is known that NTBI also enters cardiomyocytes via L and T-type calcium channels.

To minimise carryover of Fe55, cells were washed with PBS 3 times before addition of efflux medium. Efflux measurements were carried out in three independent experiments, with each experiment containing three biological replicates (3 wells of cardiomyocytes) for each genotype and condition. The range between values obtained from three biological replicates constituted between 0.07and 0.21 of the value of mean. Thus, the results were sufficiently accurate to make meaningful comparisons between genotypes and treatment conditions within each experiment, and the outcome of those comparisons was reproducible between experiments.

The reviewer raises an interesting question relating to the levels of iron export being higher in Slc40a1 C326Y knock-in cardiomyocytes than in Hamp knockout cardiomyocytes (p=0.014). We too were initially surprised at this result. One possible explanation is that FPN levels in Slc40a1 C326Y knock-in cardiomyocytes are higher than in Hamp KOs cardiomyocytes, due to the fact that FPN in the former setting is completely insensitive to any hepcidin, while FPN in the latter may still be subject to some regulation by hepcidin from non-cardiomyocyte sources.

8) Figure 1, panel A – this appears to be mRNA expression but this is not stated. If it is, how was it normalized? In panel B, as the authors pointed out, it is quite surprising that there appears to be most anti-Hamp staining under iron deficient conditions. If this is truly physiologically important staining (see point 2 above) then it becomes even more important to understand how Hamp expression is regulated (see point 3 above). Hamp is typically measured extracellularly, rather than intracellularly because it is a secreted protein; a defect in pre-protein processing might help explain the discrepancy between panels B and C in this figure. Panel D is particularly important – how many times was this experiment replicated? How reliable is the assay kit (the methods indicate that it is from "uscnk" but I don't know what that is – please specify), and is it specific for Hamp1 rather than Hamp2?

In Figure 1, panel A depicts mRNA expression of hepcidin. We have amended the corresponding figure legend to make this clear. Expression was measured using Taqman multiplex gene expression assays, where hepcidin and the house keeping gene (b-actin) were measured in the same sample. A δ CT (CT for hepcidin-CT for b-actin) value was generated for each sample. Subsequently δ CT values for each sample were normalised to the average δ CT of the control samples (hearts under control diet), to obtain a δ CT value. Relative expression was then calculated as 2-δ δ CT. The reviewer’s questions relating to the discrepancy between hepcidin transcript and protein, the possible role of protein processing and the measurement of hepcidin in supernatants are addressed in responses to points 1 and 2 above.

9) The heart sections in Figure 2B are suboptimal. It is particularly difficult to discern anatomy in the fl/fl section. In Figure 2D, the control cardiomyocytes were much smaller than those reported in Figure 1G of Xu et al., 2015, and the difference in cardiomyocyte surface area between controls and mutants was much larger. Do the authors have any idea what this difference might be due to?

We provide better heart sections in Figure 2bB In terms of cardiomyocyte size, differences in reported cardiomyocyte size between our data (range between 25 and 37 um2) and those of reported by Xu et al4 (range between 45 and 60 um2) in control animals may be due to any number of reasons including: a) the manual nature of quantitation of cardiomyocyte size, which requires drawing around individual WGA-stained cardiomyocytes, which is likely to vary between individual users, b) the method for tissue fixing and processing of the cardiac tissue, c)the area of the heart selected for histology and d) orientation of section plane relative to cardiomyocyte axis.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

The reviewers generally did not think that the authors use of tissue immunostaining to quantitate a secreted protein is bullet-proof but accept that the authors have enough additional overlapping evidence to give more confidence that the main conclusions are right. They have put together a strong case for hepcidin and ferroportin interacting in an autocrine/paracrine manner in the heart. The genetic argument from the equivalent pathology and similar survival of cardiomyocyte FpnC326Y (cardiomyocyte hepcidin resistance) and cardiomyocyte hepcidin KO is particularly appealing. The autocrine/paracrine function of hepcidin and ferroportin is an old conjecture but this is the first evidence for it.

However, the following is required for acceptance:

1) In the rebuttal there is a statement that the reviewers did not agree with. The cardiac-specific Tfr1 KO mice are not embryonic lethal – they live into the second postnatal week. The authors should be aware of this and modify the text if needed.

We thank the reviewers for drawing our attention to the erroneous statement that “cardiacTfR1 knockout are embryonic lethal”. In the original response letter, the phenotype of these mice was discussed in relation to our results showing the levels of ETC complexes as detected by the OXPHOS assay (Figure 4—figure supplement 1). As these data have now been removed from the manuscript, so has the erroneous statement in the revised response letter.

2) One reviewer did not believe that the authors had fully explained the outstanding issues with their OXPHOS blot – they thought it was poor quality with much more animal to animal variability than there should be. If possible, the authors should repeat this blot in order to show more consistency between replicates. Alternatively, the authors should delete this western.

In relation to the OXPHOS blot showing the levels of ETC complexes (Figure 4—figure supplement 1), we have now run the blot a second time. However, the variability between some of the samples persists. As such, we have opted to remove the blot from the manuscript, and have also removed all text relating to it in the Results, Discussion, Methods and figure legends section. Accordingly, we have also removed any text in our original response letter relating to this particular dataset.

https://doi.org/10.7554/eLife.19804.030

Article and author information

Author details

  1. Samira Lakhal-Littleton

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    SL-L, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    samira.lakhal-littleton@dpag.ox.ac.uk
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7797-1567
  2. Magda Wolna

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    MW, Acquisition of data
    Competing interests
    The authors declare that no competing interests exist.
  3. Yu Jin Chung

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    YJC, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  4. Helen C Christian

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    HCC, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  5. Lisa C Heather

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    LCH, Analysis and interpretation of data, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  6. Marcella Brescia

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    MB, Acquisition of data, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  7. Vicky Ball

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    VB, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  8. Rebeca Diaz

    Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    RD, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  9. Ana Santos

    Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    AS, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  10. Daniel Biggs

    Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    DB, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  11. Kieran Clarke

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    KC, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  12. Benjamin Davies

    Wellcome Trust Centre for Human Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    BD, Conception and design, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3623-600X
  13. Peter A Robbins

    Department of Physiology, Anatomy and Genetics, University of Oxford, Oxford, United Kingdom
    Contribution
    PAR, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4975-0609

Funding

British Heart Foundation (FS/12/63/29895)

  • Samira Lakhal-Littleton

Wellcome Trust (090532/Z/09/Z)

  • Samira Lakhal-Littleton

Vifor Pharma (R19874-CN004)

  • Magda Wolna

The funders funded the salary and research expenses relating to this work for two of the authors, as described above.

Acknowledgements

We thank Yixing Wu, MRC Functional Genomic Unit, University of Oxford for his help with preparation of reagents for ETC complex assays.

We thank Prof Keith Dorrington, Dr Matthew Frise (Department of Physiology, Anatomy and Genetics, University of Oxford) and Dr Edward Littleton (University Hospitals Birmingham NHS Foundation Trust, Queen Elizabeth Hospital), for critical reading of the paper.

Ethics

Animal experimentation: All animal procedures were compliant with and approved under the UK Home Office Animals (Scientific Procedures) Act 1986 (Project Licence number 30/3182). In vivo assessment of cardiac function was carried out under anesthesia (2% Isofluorane) to minimize suffering.

Reviewing Editor

  1. Richard P Harvey, The Victor Chang Cardiac Research Institute, Australia

Publication history

  1. Received: July 19, 2016
  2. Accepted: November 24, 2016
  3. Accepted Manuscript published: November 29, 2016 (version 1)
  4. Version of Record published: December 21, 2016 (version 2)

Copyright

© 2016, Lakhal-Littleton et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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