Anemic hypoxemia reduces myoblast proliferation and muscle growth in late-gestation fetal sheep

Abstract

Fetal skeletal muscle growth requires myoblast proliferation, differentiation, and fusion into myofibers in addition to protein accretion for fiber hypertrophy. Oxygen is an important regulator of this process. Therefore, we hypothesized that fetal anemic hypoxemia would inhibit skeletal muscle growth. Studies were performed in late-gestation fetal sheep that were bled to anemic and therefore hypoxemic conditions beginning at ∼125 days of gestation (term = 148 days) for 9 ± 0 days (n = 19) and compared with control fetuses (n = 16). A metabolic study was performed on gestational day ∼134 to measure fetal protein kinetic rates. Myoblast proliferation and myofiber area were determined in biceps femoris (BF), tibialis anterior (TA), and flexor digitorum superficialis (FDS) muscles. mRNA expression of muscle regulatory factors was determined in BF. Fetal arterial hematocrit and oxygen content were 28% and 52% lower, respectively, in anemic fetuses. Fetal weight and whole body protein synthesis, breakdown, and accretion rates were not different between groups. Hindlimb length, however, was 7% shorter in anemic fetuses. TA and FDS muscles weighed less, and FDS myofiber area was smaller in anemic fetuses compared with controls. The percentage of Pax7⁺ myoblasts that expressed Ki67 was lower in BF and tended to be lower in FDS from anemic fetuses indicating reduced myoblast proliferation. There was less MYOD and MYF6 mRNA expression in anemic versus control BF consistent with reduced myoblast differentiation. These results indicate that fetal anemic hypoxemia reduced muscle growth. We speculate that fetal muscle growth may be improved by strategies that increase oxygen availability.

INTRODUCTION

Birth weight is strongly associated with skeletal muscle mass in adulthood, indicating that growth during the fetal period impacts the trajectory of muscle accrual throughout the lifespan (1). The intrauterine growth restricted (IUGR) human fetus is born with lower muscle mass than their appropriately grown counterparts at a similar gestational age (2–4). Individuals who were formerly IUGR have lower muscle mass in childhood and adulthood, which has been shown to impair both their strength and metabolic health (5–10). In a well-established sheep model of severe placental insufficiency, late-gestation IUGR fetal lambs have 40% lower skeletal muscle mass when normalized to either brain weight, body weight, or hindlimb length compared with normally growing control fetuses (11). Myoblast proliferation was ∼25% lower, rates of myogenesis were ∼60% slower, and myofiber areas were reduced by ∼30% for both type I and type II fibers (11–13). Several pathophysiological factors that are characteristic of the IUGR fetus, including lower blood oxygen concentrations, lower circulating insulin and insulin-like growth factor 1 (IGF-1) concentrations, and higher catecholamine concentrations (14), are likely to have a negative impact on protein accretion rates, skeletal muscle growth, and myofiber development. In particular, fetal hypoxemia is a cardinal feature of ischemic placental disease, which includes IUGR as well as preeclampsia and chronic placental abruption (15). Chronic fetal anemia, which can result from hemolysis from maternal alloimmunization, twin anemia polycythemia sequence, placental/fetal tumors, or fetomaternal hemorrhage, will also reduce fetal O₂ content (16). These obstetrical complications carry a disproportionately high risk of perinatal morbidity and mortality, in addition to lifelong growth disturbances in the offspring.

We developed a model of late-gestation fetal anemic hypoxemia to determine the effects of lower circulating oxygen concentrations on fetal organ function. In this model, blood oxygen content in the fetus was reduced by 50%, mimicking those concentrations seen in the IUGR fetus (11) by daily blood removal from a catheterized fetal lamb from 125 to 134 days of gestation (term 148 days gestation, dGA) (17–19). In our previous work using this model, we discovered that fetuses with anemic hypoxemia compared with controls had shorter hindlimbs despite similar fetal weight (17). Skeletal muscle comprises 40% of hindlimb weight (11) and is the most metabolically active tissue within the hindlimb (20), raising the possibility that low circulating oxygen negatively impacts the growth of skeletal muscle in the fetus. In support of this, prior work in postnatal animal models and satellite cell cultures have collectively demonstrated that hypoxia negatively regulates myogenic proliferation, differentiation, and fusion into mature myofibers, otherwise referred to as myogenesis (21). Despite what is known about postnatal muscle growth and function, our understanding of the effects of hypoxemia on myogenesis during late-gestation fetal development in vivo is more limited.

Our goal was to test the effect of fetal anemic hypoxemia on amino acid metabolism and skeletal muscle growth with the hypothesis that a prolonged (9 day) exposure to anemic hypoxemia would reduce myoblast proliferation and hypertrophy, thus leading to smaller muscle mass. Muscle is a major site of protein turnover and plays an important role in whole-body protein metabolism by serving as a reservoir for amino acids (22). Thus, we measured umbilical (net fetal) amino acid uptake rates and whole body protein synthesis and accretion rates in response to fetal anemic hypoxemia. We determined weights, myofiber cross-sectional area, myosin heavy chain (MHC) expression to determine relative fiber type proportions (type I vs. type IIa MHC), and myoblast proliferative capacity in three different skeletal muscles within the fetal hindlimb that develop relatively more (tibialis anterior, flexor digitorum superficialis) versus less (biceps femoris) type I oxidative fibers in adult sheep (23, 24). Finally, we measured the expression of key genes and proteins within pathways that regulate muscle differentiation, protein breakdown, and protein synthesis.

MATERIALS AND METHODS

Fetal Sheep Model of Anemic Hypoxemia

Studies were conducted on pregnant Columbia-Rambouillet sheep carrying a singleton fetus at the Perinatal Research Center in Aurora, Colorado with approval of the Institutional Animal Care and Use Committee, University of Colorado School of Medicine. This center is accredited by AAALAC International. Sheep were fasted for 24 h and thirsted for 12 h before surgery. In 35 pregnant sheep at 119 ± 0 dGA, fetal catheters were surgically placed into the descending aorta and femoral vein via pedal incisions and into the common umbilical vein via placement of a nonocclusive catheter into one of the umbilical veins (18). An amniotic catheter was placed. Maternal catheters were placed into the femoral artery and vein via a groin incision (18). Surgery was performed with anesthesia and pain control as follows. A maternal jugular venous catheter was placed for administration of diazepam (0.2 mg·kg⁻¹) and ketamine (20 mg·kg⁻¹) and ewes were maintained on isoflurane inhalation anesthesia (2%–4%) for the remainder of the surgical procedure. Postsurgical pain control was achieved with flunixin meglumine (2.2 mg·kg⁻¹ per day intramuscularly) on the day of the operation and the following 1–3 days based on assessment of the animal’s comfort. Immediately before surgery, the ewe also received procaine penicillin G (600,000 units, intramuscularly) and ampicillin (500 mg) was instilled into the amniotic cavity before closure of the hysterotomy and then three times a week postoperatively via an amniotic catheter.

Beginning at least 5 days after surgery on 125 ± 0 dGA, fetuses were randomly assigned to be bled with isovolumetric replacement by 0.9% NaCl to anemic conditions for 9 ± 0 days (anemic, n = 19 fetuses) or not bled but otherwise treated the same as the anemic fetuses with daily monitoring of blood gases and hematocrit (control, n = 16 fetuses). The target arterial blood O₂ content for the anemic fetuses was 2.0 mmol·L⁻¹, which is the mean arterial blood O₂ content in IUGR fetal sheep with placental insufficiency (25, 26). The amount of blood removed daily was determined using a previously established formula taking into account fetal hematocrit and the target arterial blood O₂ content (19). The impact of fetal anemic hypoxemia on fetal arterial biochemistry, glucose production, and glucose-stimulated insulin secretion in these fetuses has been previously published (17, 18).

The first set of 18 fetuses in this study received a metabolic study at the end of the 9-day experimental period on 134 ± 0 dGA to determine umbilical blood flow and amino acid, glucose, lactate, and oxygen uptake rates (anemic, n = 11 fetuses; control, n = 7 fetuses). Because of catheter failure, complete metabolic studies were completed on seven animals in each group. One anemic fetus did not survive to necropsy following the metabolic study, but there were no other unexpected fetal deaths. A shorter fetal hindlimb length was observed in anemic fetuses (17); therefore, the second set of 17 fetuses in this study (anemic, n = 8 fetuses; control, n = 9 fetuses) did not receive a metabolic study but instead received a leucine-stimulated insulin secretion study for which methods and results are published (17) and muscles were collected for weight and histology measurements.

In Vivo Metabolic Studies

After samples were collected at time 0 for naturally occurring isotopic enrichments, a solution containing ethanol (300 mg bolus followed by 12.8 mg·min⁻¹; AAPER Alcohol and Chemical, Shelbyville, KY) was infused into the fetal venous circulation to measure umbilical blood flow using the transplacental diffusion method (27). Fetuses also received a primed (30 μmol·kg⁻¹) intravenous infusion with l-[1-¹³C]leucine (0.5 μmol·min·kg⁻¹ estimated fetal weight; Cambridge Isotope Laboratories, Woburn, MA) to measure fetal leucine metabolism (28). After 180 min of tracer infusion equilibration, four fetal blood draws were obtained 20 min apart to represent steady-state conditions for measurements of fetal arterial and umbilical venous plasma glucose, lactate, and amino acid concentrations including α-ketoisocaproic acid (KIC) and whole blood O₂ content. Isotopic enrichments of [1-¹³C]leucine and KIC in fetal plasma and ¹³CO₂ in fetal whole blood were measured under steady-state conditions. Immediately following the metabolic study, glucose- and arginine-stimulated insulin secretion studies were performed, for which methods and results have been previously published (17). During the metabolic study period, the fetal blood removed was replaced with heparinized maternal whole blood diluted with 0.9% NaCl to match the hematocrit of the fetal blood.

Umbilical blood flow rates were calculated by dividing umbilical plasma flow by one minus fractional fetal hematocrit. Umbilical venous-fetal arterial differences in plasma amino acids, glucose, and lactate concentrations were multiplied by the umbilical plasma flow rate (Fick principle) to calculate umbilical (net fetal) substrate uptake rates. Net fetal oxygen consumption rates were calculated by multiplying umbilical venous-fetal arterial differences in whole blood O₂ content by umbilical blood flow rate. The net fetal nitrogen uptake rate was determined by multiplying the number of nitrogen molecules in each amino acid by their respective net fetal uptake rates and adding them together. [1-¹³C]leucine tracer fluxes between the placenta and fetal plasma and between fetal plasma and fetal tissues were calculated as previously described (28, 29).

Biochemical Analysis

Fetal and maternal arterial plasma glucose and lactate concentrations were measured using a Yellow Springs Instrument 2700 (Yellow Springs Instruments, Yellow Springs, OH). Blood hematocrit, pH, partial pressure of O₂ ($\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$), partial pressure of carbon dioxide ($\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$), and hemoglobin-O₂ saturation were measured with a Blood Gas Analyzer ABL 825 (Radiometer, Copenhagen, Denmark). O₂ content of the blood was calculated by the ABL 825 analyzer. Arterial plasma insulin, IGF-1, and cortisol were measured by an enzyme-linked immunosorbent assay (ELISA; Insulin: ALPCO, Windham, NH; intra- and interassay coefficients of variation = 5.6% and 4.7%, respectively; sensitivity = 0.14 ng·mL⁻¹; IGF-1: ALPCO; intra- and interassay coefficients of variation, 3.1% and 5.6%, respectively; sensitivity, 0.09 ng·mL⁻¹; cortisol: ALPCO; intra- and interassay coefficients of variation = 4.6% and 5.8%, respectively; sensitivity = 1.0 ng·mL⁻¹) and norepinephrine by high-performance liquid chromatography (HPLC; Model 2475, Waters; intra- and interassay coefficients of variation = 9.2% and 9.0%, respectively; sensitivity = 170 pg·mL⁻¹) (30). Amino acid concentrations were measured using a Dionex 300 model 4500 amino acid analyzer (Dionex, Sunnyvale, CA). Isotopic enrichments of [1-¹³C]leucine and [1-¹³C]KIC were determined as previously described (6890 N/5975 Inert XL MSD mass spectrometer, Agilent Technologies, Inc.) (11). Isotopic enrichments of ¹³CO₂/¹²CO₂ in fetal blood were measured using isotope ratio mass spectrometry as previously described (Delta V Advantage, Thermo Electron North America, LLC) (11).

Necropsy

The day following all in vivo studies, pregnant sheep were sedated with diazepam (0.2 mg·kg⁻¹) and ketamine (20 mg·kg⁻¹) intravenously and fetuses were delivered via maternal laparotomy and hysterotomy. The biceps femoris (BF) muscle was exposed and a biopsy was obtained from the anesthetized fetus and immediately frozen in liquid nitrogen and stored at −80°C for the purpose of RNA and protein analysis. Fetal hindlimb length was measured from the femoral head to the top edge of the hoof. Because we discovered a reduction in hindlimb length in the first set of anemic fetuses compared with controls [33.8 ± 0.7 control (n = 7 fetuses), 31.8 ± 0.5 cm anemic (n = 10 fetuses); P < 0.05], fetal hindlimb muscles including the BF, tibialis anterior (TA), flexor digitorum superficialis (FDS), and gastrocnemius were collected within 15 min postmortem and weighed to further evaluate the effects of anemia on skeletal muscle-specific growth in the second set of fetuses (anemic, n = 8 fetuses; control, n = 9 fetuses). Muscle mid-bellies from the BF, TA, and FDS muscles were placed on corkboard thinly coated with optimal cutting temperature media, frozen in liquid nitrogen-cooled isopentane for 60 s, and stored at −80°C. The mother and fetus were killed with pentobarbital sodium (390 mg·mL⁻¹ intravenously; 12 and 2 mL, respectively; Bortech Pharmaceuticals, Dearborn, MI).

Histology of Fetal Skeletal Muscles

Isopentane-preserved muscles were removed from corkboard and 14-µm sections were prepared on Superfrost Plus Microscope slides (Fisher Scientific, Pittsburgh, PA) using a Cryostat (Leica, Buffalo Grove, IL; anemic, n = 8 fetuses; control, n = 9 fetuses). Cryosections were air-dried for 30 min and fixed with 4% paraformaldehyde for 5 min (for Pax7 staining only) at room temperature. Slides were briefly washed in phosphate-buffered saline (PBS) and blocked with 5% normal goat serum (NGS) that included anti-laminin rabbit polyclonal IgG (1:500; Sigma Aldrich, St. Louis, MO; Cat. No. L9393; RRID: AB_477163) in permeabilizing/blocking solution (PBS: 0.1% Triton-X-100, 0.12% bovine serum albumin, 0.12% nonfat dry milk in PBS) for 1 h. After a brief wash in PBS, muscle sections were incubated with the primary antibody of interest in PBS and 5% NGS. These included anti-PAX7 mouse monoclonal IgG (1:100; Developed by Tokyo Institute of Technology and obtained from the Developmental Studies Hybridoma Bank, created by the National Institute of Child Health and Human Development (NICHD) of the National Institutes of Health (NIH) and maintained at the Depart ment of Biology, University of Iowa, Iowa City, IA; Cat. No. PAX7-c; RRID: AB_2299243), anti-Ki67 rabbit monoclonal IgG (1:250; Cell Signaling Technology, Danvers, MA; Cat. No. 9129; RRID: AB_2687446), anti-MHC type I (slow) mouse monoclonal IgG (1:1,000; Developmental Studies Hybridoma Bank; Cat. No. BA-D5; RRID: AB_2235587), and anti-MHC type IIa mouse IgG (1:2 incubated overnight at 4°C; a gift from Dr. Leslie Leinwand, Boulder, CO; RRID: AB_2147165). Slides were washed in PBS with gentle shaking for 20 min three times. Immunocomplexes were detected with affinity-purified IgG antiserum conjugated to Alexa Fluor 594 (1:500; Thermo Fisher Scientific, Waltham, MA; Cat. No. A11005; RRID: AB_2534073), Alexa Fluor 488 (1:500; Thermo Fisher Scientific, Cat. No. A11008; RRID: AB_143165), Cy2 (1:500; Jackson ImmunoResearch Laboratories, Inc., Bar Harbor, ME; Cat. No. 175–225-150; RRID: AB_2340826) in PBS and 5% NGS for 1 h at 37°C. Slides were washed briefly in PBS, counterstained with DAPI (1:1,000) and mounted with Fluoromount G (Thermo Fisher Scientific, Rockford, IL). Fluorescent images were visualized on an Olympus IX83 Motorized Fluorescent Microscope with Dual Sensor DP80 Camera to capture images. Images were quantified using CellSens Dimension Imaging Software (Olympus). Pax7⁺ and Ki67⁺ nuclei were expressed as a percentage of total nuclei identified with DAPI. Dual-stained Ki67⁺ and Pax7⁺ nuclei were expressed as a percentage of total Pax7⁺ nuclei. Myofibers that expressed either type I or IIa MHC were expressed as a percentage of total myofibers identified and the cross-sectional areas of type I and IIa MHC myofibers were determined. A minimum of 2,600 nuclei or 1,000 myofibers were analyzed from a minimum of four individual muscle sections for each muscle type.

RNA Analysis

RNA was extracted from pulverized −80°C BF (100 mg) and reverse transcribed into complimentary DNA (cDNA) as previously described (anemic, n = 18 fetuses; control, n = 15–16 fetuses) (31). Extraction was by immersion in TRIzol (Invitrogen, Thermo Fisher Scientific) and homogenization. To separate nucleic acids and proteins, the homogenate was mixed with chloroform and centrifuged at 15,000 rpm at 4°C. The aqueous phase was then removed and an RNeasy mini column (Qiagen, Hilden, Germany) was used to isolate total RNA. Purity and quantity of RNA were determined with a spectrophotometer (Nanodrop 1000). All samples were considered to be pure based on an A260/A280 absorbance ratio of ∼2.1 (range 2.07–2.13). RNA integrity was verified based on visible bands for 5S, 18S, and 28S using electrophoresis. Two micrograms of total RNA were reverse transcribed using SuperScript III First-Strand Synthesis SuperMix (Invitrogen) and Oligo dT 18–20 (Invitrogen) at 50°C for 1 h.

Quantitative PCR assays on a 1:10 dilution (with sterile water) of the reverse transcription cDNA for GAPDH, ACTA1, RPS15, MYF5, MYOG, MYOD1, MYF6, PIKC3C, MAP1LC3A, BNIP3, CTSL, BECN1, ATF4, TRIM63, and FBXO32 were performed and validated as previously described (31, 32). The total volume of each reaction was 10 µL: 4 µL cDNA, 0.5 µL for each primer (0.5 mol·L⁻¹), and 5 µL Faststart Universal SYBR Green MM (Roche, Basel, Switzerland). The temperature cycles were 5 min at 95°C to activate the enzyme, then 40 cycles of 30 s at 60°C, 30 s at 72°C, and 10 s at 96°C. To verify the correct amplification of the reaction, we examined the melt-curve analysis to ensure a single peak. The cDNA samples were analyzed in triplicate and the standard curve method of relative quantification was utilized (33). Genes of interest were normalized to the mean of the reference genes GAPDH, RPS15, and ACTA1, which were not different between treatment groups. Results are presented relative to controls.

Western Blot Analysis

Biopsies from the BF were ground in liquid nitrogen using mortar and pestle and 100 mg of muscle was homogenized in 500 µL commercially available lysis buffer (Cell Signaling Technology) containing 1% phosphatase and 2% protease inhibitors (Sigma-Aldrich). Protein concentrations of cell lysates were quantified using Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Twenty micrograms of protein were combined with 250 mM dithiothreitol, heated to 90°C for 5 min, and loaded onto a precast NuPAGE 4%–12% Bis-Tris Mini Gel 1.0 mm × 17 well (Thermo Fisher Scientific). Electrophoresis was performed, and protein was transferred to nitrocellulose membranes blocked with LI-COR Odyssey Blocking Buffer (LI-COR Biosciences, Lincoln, NE) at a 1:1 ratio with tris-buffered saline (TBS). All primary antibodies were diluted in LI-COR Odyssey Blocking buffer in TBS with 0.1% tween. The membrane was serially incubated overnight at 4°C in antibodies (all from Cell Signaling Technology) against phosphorylated Akt (S473; Cat. No. 9271; RRID: AB_329826 at 1:1,000), total Akt (Cat. No. 2920; RRID: AB_1147620 at 1:1,000), phosphorylated 4E-BP1 (T37/46; Cat No. 9459S; RRID: AB_330985 at 1:1,000), total 4E-BP1 (Cat. No. 9452; RRID: AB_331692 at 1:1,000), phosphorylated AMPK α (T172; Cat. No. 2531 L; RRID: AB_330330 at 1:1,000), and total AMPK α (Cat. No. 2793S; RRID: AB_915794 at 1:1,000). β-Actin (Cat. No. 3700S; RRID: AB_2242334 at 1:1,000) was used to account for differences in loading and transfer efficiency. Bands were visualized and quantified using near-infrared fluorescence exposed on the Odyssey FC-0788 (LI-COR Biosciences) using either IRDye 800CW goat anti-rabbit (LI-COR; Cat. No. 926–32211; RRID: AB_621843 at 1:5,000) or IRDye 700CW goat anti-mouse (LI-COR; Cat. No. 926–68070; RRID: AB_10956588 at 1:5,000) diluted in LI-COR Odyssey Blocking Buffer at a 1:1 ratio with TBS.

Statistical Analysis

Statistical analysis was performed using SAS version 9.2 (SAS Institute), GraphPad Prism version 6.0, and Microsoft Excel 2016. Results are expressed as means ± standard error (SE). Comparisons between control and anemic fetuses were made by Student’s t test or Mann–Whitney test for parametric and nonparametric data, respectively. P values ≤0.05 were accepted as reaching statistical significance and P values <0.1 were considered as trending toward significance. After determining that fetal sex was not significant for a given outcome in a preliminary two-way ANOVA (male or female, anemic or control, and the interaction), we pooled data from male and female fetuses. If fetal sex or the interaction was significant (P < 0.05), we included the term for fetal sex and presented the summary statistics. There was a significant effect of fetal sex on fetal weight and MYOD mRNA expression in the BF. The interaction of fetal sex with group (anemic or control) was significant for the mRNA expression of MAP1LC3A and BNIP3 mRNA in the BF.

RESULTS

Maternal and Fetal Biochemical Measurements, Substrate, and Hormone Concentrations

As previously reported (17), maternal values for blood pH, $\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$, $\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$, hemoglobin-O₂ saturation ($\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$) O₂ content, hematocrit, and plasma glucose and lactate concentrations were similar between groups at the end of the experimental period (data not shown). Fetal arterial blood biochemical parameters on the day of the metabolic study (at the end of the anemic or control treatment) are reported in Table 1, some of which have been previously published (17). Fetal pH was not different between groups, and fetal blood $\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$ was higher in anemic fetuses compared with control fetuses (P < 0.05). Fetal $\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$ (P < 0.01), $\mathrm{S}{{\mathrm{a}}_{\mathrm{O}}}_2$ (P < 0.0001), and O₂ content (P < 0.0001) were 14%, 28%, and 50% lower, respectively, in anemic fetuses compared with controls. Fetal plasma glucose was 30% lower in anemic fetuses compared with controls (P < 0.01). Most fetal arterial amino acid concentrations were similar between groups except for lower concentrations of asparagine, aspartate, glutamate, and proline in anemic fetuses (P < 0.05). IGF-1 concentrations were 28% lower and norepinephrine concentrations were 80% higher in anemic fetuses compared with controls (P < 0.01).

Table 1.

Fetal arterial biochemistry at the end of the study

	Control	Anemic
Blood gas measurements
pH	7.35 ± 0.00	7.34 ± 0.00
$\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$, mmHg	51.2 ± 0.6	53.2 ± 0.6*
$\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$, mmHg	21.3 ± 0.6	18.3 ± 0.5**
$\mathrm{S}{{\mathrm{a}}_{\mathrm{O}}}_2$, %	51.7 ± 2.1	37.4 ± 2.1****
O2 content, mmol·L−1	3.57 ± 0.12	1.72 ± 0.12****
Hct, %	35.5 ± 0.9	23.5 ± 1.0****
Plasma substrates
Glucose, mmol·L−1	1.0 ± 0.1	1.3 ± 0.1**
Lactate, mmol·L−1	2.1 ± 0.1	2.4 ± 0.1
Alanine, nmol·L−1	293 ± 18	321 ± 16
Arginine, nmol·L−1	100 ± 7	87 ± 8
Asparagine, nmol·L−1	43 ± 2	36 ± 3*
Aspartate, nmol·L−1	18 ± 1	12 ± 1**
Cysteine, nmol·L−1	11 ± 1	10 ± 1
Glutamate, nmol·L−1	37 ± 3	24 ± 3**
Glutamine, nmol·L−1	379 ± 14	367 ± 16
Glycine, nmol·L−1	344 ± 21	291 ± 20
Histidine, nmol·L−1	53 ± 2	52 ± 3
Isoleucine, nmol·L−1	85 ± 5	98 ± 5
Leucine, nmol·L−1	121 ± 5	120 ± 6
Lysine, nmol·L−1	76 ± 4	65 ± 4
Methionine, nmol·L−1	78 ± 3	68 ± 4
Phenylalanine, nmol·L−1	93 ± 5	90 ± 8
Proline, nmol·L−1	143 ± 5	127 ± 5*
Serine, nmol·L−1	685 ± 46	638 ± 39
Taurine, nmol·L−1	69 ± 13	74 ± 9
Threonine, nmol·L−1	293 ± 29	275 ± 39
Tryptophan, nmol·L−1	37 ± 1	39 ± 2
Tyrosine, nmol·L−1	99 ± 8	90 ± 6
Valine, nmol·L−1#	326 ± 8	350 ± 17
Plasma hormones
Insulin, ng·mL−1	0.33 ± 0.04	0.33 ± 0.03
-1, ng·mL−1	123.5 ± 8.8	88.4 ± 5.6**
Glucagon, pg·mL−1#	39.1 ± 4.4	64.6 ± 7.9**
Cortisol, ng·mL−1	10.6 ± 1.7	16.6 ± 2.4†
Norepinephrine, pg·mL−1	528.6 ± 97.8	952.7 ± 95.3**

Values are expressed as means ± SE. Control, n = 14–15 fetuses; anemic, n = 16–19 fetuses.

*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, respectively by Student’s t test or the Mann–Whitney test (#). †P < 0.06 by Student’s t test. IGF-1, insulin-like growth factor-1; $\text{P}{\text{a}}_{\text{CO}}{}_{{}_2}$, partial pressure of carbon dioxide; $\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$, partial pressure of oxygen; $\mathrm{S}{{\mathrm{a}}_{\mathrm{O}}}_2$, hemoglobin-oxygen saturation.

Net Umbilical Substrate Uptake Rates, Oxygen Consumption, and Fetal Protein Metabolism

Umbilical blood flow and uptake rates of glucose and lactate were similar between groups and have been previously published (Table 2) (18). Umbilical oxygen uptake rates were 16% lower in anemic fetuses compared with controls (P < 0.05) (18). There were no differences between the groups for uptake rates of any of the amino acids; total carbon from amino acids; total carbon from amino acids, glucose, and lactate; and total nitrogen. There were no differences for leucine flux rates among placenta, fetus, and fetal tissues. Fetal protein synthesis and accretion rates were similar between groups.

Table 2.

Umbilical blood flow, substrate uptake, and leucine flux rates

	Control	Anemic
Umbilical blood flows, mL·min−1·kg−1
Blood flow	156.1 ± 14.3	138.8 ± 11.0
Plasma flow	103.7 ± 10.7	105.9 ± 9.5
Umbilical substrate uptake rates, µmol·min−1·kg−1
Glucose	30.2 ± 1.9	29.5 ± 2.7
Lactate	28.0 ± 2.4	23.5 ± 2.9
Oxygen	321 ± 15	270 ± 17*
Alanine	2.6 ± 0.4	2.8 ± 0.5
Arginine	2.2 ± 0.3	1.9 ± 0.2
Asparagine	1.0 ± 0.2	1.1 ± 0.3
Aspartate	−0.1 ± 0.1	0.1 ± 0.1
Cysteine#	0.4 ± 0.1	0.2 ± 0.0
Glutamate	−2.6 ± 0.4	−1.8 ± 0.3
Glutamine#	4.8 ± 0.8	4.2 ± 0.3
Glycine	2.5 ± 0.5	2.2 ± 0.3
Histidine	0.6 ± 0.1	0.5 ± 0.1
Isoleucine	1.6 ± 0.1	1.9 ± 0.1
Leucine	2.6 ± 0.2	2.6 ± 0.3
Lysine	2.0 ± 0.2	1.5 ± 0.1
Methionine	0.8 ± 0.1	0.8 ± 0.1
Phenylalanine	1.3 ± 0.2	1.1 ± 0.2
Proline	2.0 ± 0.3	1.7 ± 0.3
Serine	−1.7 ± 0.6	−0.2 ± 0.5
Taurine	0.1 ± 0.1	0.1 ± 0.1
Threonine	1.8 ± 0.4	1.7 ± 0.3
Tryptophan	0.4 ± 0.1	0.4 ± 0.0
Tyrosine	1.1 ± 0.1	1.1 ± 0.1
Valine	3.3 ± 0.5	3.5 ± 0.5
Total amino acid carbon	139.6 ± 20.2	141.5 ± 14.5
Total amino acid, glucose, and lactate carbon	424.4 ± 27.3	408.2 ± 29.6
Total amino acid nitrogen	43.4 ± 6.9	42.9 ± 4.1
Leucine flux rates, µmol·min−1·kg−1
Disposal rate	8.3 ± 0.3	8.3 ± 0.3
Flux into fetal blood from placenta	4.2 ± 0.3	3.6 ± 0.3
Flux into placenta from fetal blood	1.6 ± 0.2	0.9 ± 0.3
Flux into fetal tissues from fetal blood#	6.7 ± 0.1	7.4 ± 0.5
Oxidation rate	2.0 ± 0.1	1.9 ± 0.1
Flux into fetal protein synthesis#	5.2 ± 0.1	5.9 ± 0.5
Flux into fetal blood from fetal protein breakdown#	4.1 ± 0.2	4.8 ± 0.4
Flux into net fetal protein accretion	1.1 ± 0.2	1.2 ± 0.2

Values are expressed as means ± SE. Control, n = 7 fetuses; Anemic, n = 7 fetuses.

*P < 0.05 by Student’s t test or the Mann–Whitney test (#).

Fetal Weight, Hindlimb Muscle Weights, and Myofiber Size

Male fetuses were 11% heavier than female fetuses (P < 0.005); however, there was not an interaction between fetal sex and group (anemic vs. control). Additionally, there was no impact of treatment group on fetal weight (summary statistics: 2.93 ± 0.11 kg, control females, n = 10 fetuses; 3.35 ± 0.11 kg, control males, n = 6 fetuses; 2.90 ± 0.09 kg, anemic females, n = 13 fetuses; 3.20 ± 0.10 kg, anemic males, n = 6 fetuses). Lower limb lengths were shorter in the anemic fetuses (35.1 ± 0.6 cm, controls, n = 16 fetuses vs. 32.5 ± 0.4 cm, anemics, n = 18 fetuses, P < 0.001) as previously reported (17). TA (P < 0.05) and FDS (P < 0.005) muscles from anemic fetuses weighed less compared with controls, and the summed weight of hindlimb muscles collected from anemic fetuses tended toward being lower than controls (P < 0.1; Table 3). When hindlimb muscle weights were normalized to hindlimb length, there were no differences between groups (Table 3). The proportion and cross-sectional areas of type I and type IIa fibers were determined in the BF, TA, and FDS muscles (Fig. 1). The relative proportion of the two different fiber types was similar between control and anemic fetuses (Fig. 1B). The cross-sectional areas of both type I and IIa fibers in the FDS were smaller in anemic fetuses compared with controls (P < 0.05) but not within BF and TA (Fig. 1C).

Table 3.

Fetal hindlimb muscle weights

	Control	Anemic
Biceps femoris, g	18.7 ± 0.6	17.4 ± 1.1
Relative to hindlimb length, g·cm−1	0.52 ± 0.03	0.52 ± 0.04
Gastrocnemius, g	8.8 ± 0.4	8.1 ± 0.4
Relative to hindlimb length, g·cm−1	0.27 ± 0.01	0.28 ± 0.01
Tibialis anterior, g	4.1 ± 0.1	3.4 ± 0.2*
Relative to hindlimb length, g·cm−1	0.12 ± 0.01	0.10 ± 0.01
Flexor digitorum superficialis, g	2.9 ± 0.1	2.4 ± 0.1**
Relative to hindlimb length, g·cm−1	0.08 ± 0.00	0.07 ± 0.00
Summed hindlimb muscles, g	34.5 ± 0.9	31.2 ± 1.6†
Relative to hindlimb length, g·cm−1	0.96 ± 0.04	0.93 ± 0.06

Values are expressed as means ± SE. Control, n = 9 fetuses; Anemic, n = 8 fetuses.

*P < 0.05 and **P < 0.005, respectively, by Student’s t test. †P < 0.1 by Student’s t test.

Figure 1.

Anemic fetuses have smaller flexor digitorum superficialis (FDS) myofibers. Representative images of the flexor digitorum superficialis (FDS) from a control and an anemic fetus demonstrate myosin heavy chain (MHC) distribution and myofiber size using antibodies against laminin (blue) and type I and type IIa MHC (green; A). Tissue sections from the biceps femoris (BF), tibialis anterior (TA), and FDS muscles were similarly stained in control (gray bars, n = 9 for TA and FDS, n = 8 for BF) and anemic (white bars, n = 8) fetuses and demonstrated that the percentage of fiber numbers represented by each type was not different between groups (B), and fiber type specific cross-sectional areas in the FDS, but not the BF or TA, were lower in anemic fetuses compared with controls (C). *P < 0.05 by Student’s t test. Means, individual data, and SE bars are shown.

Fetal Myoblast Proliferation

To determine the effect of fetal anemic hypoxemia on the pool of myoblasts identified by the expression of Pax7 and their proliferative capacity as determined by expression of Ki67 in fetal hindlimb skeletal muscle, the percentage of all nuclei that express Pax7, the number of Pax7⁺ nuclei per myofiber, and the proportion of Pax7⁺ nuclei positive for Ki67 were measured in the BF, TA, and FDS from control and anemic fetuses (Fig. 2). There were no differences between control and anemic fetuses for the percentage of total nuclei that were Pax7⁺ or for the ratio of Pax7⁺ myoblasts per myofiber (Fig. 2, B and C). However, the percentage of total nuclei that expressed Ki67 was 28% lower the percentage of Pax7⁺ nuclei that expressed Ki67 was 31% lower in the BF from anemic fetuses (P < 0.05) and tended toward being lower in the FDS (P < 0.1), but not the TA (Fig. 2, D and E).

Figure 2.

Myoblast proliferation is inhibited by anemic hypoxemia. Representative images of the biceps femoris (BF) from a control and an anemic fetus demonstrate proliferating myoblasts using antibodies against Pax7 (red), the proliferation marker Ki67 (green), and counter stained with a nuclear stain (DAPI, blue; A). Tissue sections from the BF, tibialis anterior (TA), and flexor digitorum superficialis (FDS) muscles were similarly stained in control (gray bars, n = 8) and anemic (white bars, n = 8) fetuses and demonstrated that the percentage of total nuclei which were Pax7⁺ (B) and the ratio of Pax7⁺ cells per myofiber (C) were not different between groups. There were fewer total nuclei that were Ki67⁺ (D) and less Pax7⁺ nuclei that were also Ki67⁺ in the BF with a trend toward fewer Ki67⁺ nuclei in the FDS (E) from anemic fetuses compare to controls, but not the TA. *P < 0.05 and †P < 0.1 by Student’s t test. Means, individual data, and SE bars are shown.

mRNA and Protein Expression

mRNA expression of the muscle regulatory factor MYOD1 was 31% lower in anemic fetuses compared with controls (P < 0.05; Table 4) and was 28% higher in females than males (P < 0.05). However, there was not an interaction between fetal sex and group (summary statistics: 1.16 ± 0.15 ratio to reference genes, control females, n = 9 fetuses; 0.71 ± 0.05 ratio, anemic females, n = 12 fetuses; 0.76 ± 0.07 ratio, control males, n = 6 fetuses; 0.64 ± 0.08 ratio, anemic males, n = 6 fetuses; P < 0.05). MYF6 mRNA expression was 18% lower in anemic fetuses compared with controls (P < 0.05) but expression of the other muscle regulatory factors MYF5 and MYOG were not statistically different between groups (Table 4). mRNA expression of regulators of protein breakdown were similar (CTSL, ATF4) or lower (PIKC3C, BECN1, TRIM63, FBXO32) in anemic fetuses compared with controls (Table 4). There was a significant interaction effect between fetal sex and group (anemic vs. control) for the mRNA expression of two regulators of autophagy, MAP1LC3A and BNIP3 in the BF (P < 0.05). For MAP1LC3A mRNA expression was lower in anemic females compared with control females (1.08 ± 0.08 ratio, control females, n = 9 fetuses vs. 0.82 ± 0.04 ratio, anemic females, n = 12 fetuses, P < 0.01). There was no difference between control and anemic males (0.88 ± 0.08 ratio, control males, n = 6 fetuses vs. 0.94 ± 0.09 ratio, anemic males, n = 6 fetuses). For BNIP3 mRNA expression, there were no statistically significant differences in the individual means comparisons (1.09 ± 0.09 ratio, control females, n = 9 fetuses; 0.99 ± 0.05 ratio, anemic females, n = 12 fetuses; 0.86 ± 0.08 ratio, control males, n = 6 fetuses; 1.14 ± 0.17 ratio, anemic males, n = 6 fetuses). We investigated the activation status of three signaling proteins involved in the regulation of protein synthesis (Akt, 4E-BP1, and AMPK) but found no differences in either the phosphorylated (active) or total protein concentrations between groups (Table 4).

Table 4.

Expression of mRNA and protein in the biceps femoris muscle

	Control	Anemic
mRNA expression (ratio)
Muscle regulatory factors
MYF5	1.00 ± 0.12	0.81 ± 0.06
MYOD1	1.00 ± 0.11	0.69 ± 0.04*
MYOG#	1.00 ± 0.09	0.80 ± 0.05
MYF6	1.00 ± 0.06	0.82 ± 0.04*
Regulators of autophagy
PIKC3C	1.00 ± 0.08	0.75 ± 0.05**
CTSL	1.00 ± 0.06	0.90 ± 0.05
BECN1	1.00 ± 0.06	0.77 ± 0.05**
ATF4	1.00 ± 0.06	0.93 ± 0.06
Regulators of protein ubiquitination
TRIM63	1.00 ± 0.07	0.70 ± 0.04****
FBXO32	1.00 ± 0.11	0.75 ± 0.06*
Protein expression (ratio)
Intracellular signaling proteins
Phosphorylated Akt (S473)	1.00 ± 0.11	0.99 ± 0.10
Total Akt	1.00 ± 0.04	0.91 ± 0.04
Phosphorylated 4E-BP1 (T37/46)	1.00 ± 0.03	1.03 ± 0.04
Total 4E-BP1	1.00 ± 0.15	1.19 ± 0.12
Phosphorylated AMPK (T172)	1.00 ± 0.09	1.18 ± 0.10
Total AMPK#	1.00 ± 0.13	0.95 ± 0.07

Values are expressed as means ± SE. Control, n = 15–16 fetuses; Anemic, n = 18 fetuses. *P < 0.05, **P < 0.01, ***P < 0.005, ****P < 0.001, respectively by Student’s t test or the Mann–Whitney test (#).

DISCUSSION

In a sheep model of prolonged (9-day) fetal anemic hypoxemia during late gestation with reduced blood O₂ content, reduced fetal oxygen consumption rates, and shorter hindlimb length compared with controls (17, 18), we demonstrated a reduction in hindlimb muscle weight and fewer proliferating myoblasts that was muscle specific. Muscle weights of the TA and FDS within the fetal hindlimb were lower and the myofiber cross-sectional area in the FDS muscle was lower in anemic fetuses compared with controls, indicating reduced myofiber hypertrophy. The expression levels of Ki67 within all muscle nuclei and within Pax7⁺ myoblasts were lower in the anemic BF muscle, reflecting reduced cell proliferation. We also demonstrated lower expression of the myogenic regulatory factors in the BF, further suggesting downregulation of the myogenic program. Consistent with preserved fetal weight in this model, net fetal substrate uptake rates, including for glucose, lactate, amino acids, total carbon, and total nitrogen were similar between hypoxemic and control groups, as were fetal whole-body protein kinetic rates as measured by a leucine stable isotope tracer.

Reduced fetal myoblast proliferation in response to anemic hypoxemia as demonstrated in this study is consistent with several previous studies that have shown a negative effect of hypoxia on myogenesis in adult animal models and immortalized cell lines (21, 34–38). In both L6 rat and C2C12 murine myoblasts in vitro, it has been shown that optimal myoblast proliferation and differentiation occur at 3%–6% oxygen, which is similar to the partial pressure of oxygen within tissue beds in vivo in adult humans and animals (21, 34). Hypoxic conditions at ≤1% oxygen impair proliferation in vitro (35–37). Hypoxia also has been shown to limit the differentiation of myoblasts into multinucleated myotubes in vitro, with inhibition of differentiation below 2% oxygen with maximal inhibition at 0.01% oxygen in murine C2C12 (38) and rat L6 (19) cell lines. Hypoxia markedly inhibited MyoD, Myf5, and MyoG mRNA expression in C2C12 murine myoblasts, and protein expression correlated with their respective mRNA expression (35, 38). Overexpression of MyoD in hypoxic, differentiating myoblasts restored the myogenic differentiation program (38), indicating that MyoD is one of the major regulators of the myogenic response to hypoxia. In anemic hypoxemic fetuses, we similarly demonstrated a significant reduction in MYOD and MYF6 mRNA expression but were not able to measure protein expression of the muscle regulatory factors due to lack of availability of sheep-specific antibodies. It should be noted that the response to hypoxia, at least in vitro, has not always demonstrated a reduction in myogenesis, as 1% oxygen exposure to primary cultured bovine satellite cells and 3% oxygen exposure to aged rat satellite cells increased proliferation, formation of myotubes, and expression of MyoD (39, 40). Contrasting responses among studies are likely dependent on cell type (transformed cell line vs. primary satellite cells), species of origin, level of oxygen exposure, and adaptability of the cell to hypoxic conditions (40). However, our findings of reduced myoblast proliferation and downregulation of myogenic regulatory genes in anemic hypoxemic fetuses are consistent with substantial evidence that myoblasts are sensitive to the concentration of oxygen in terms of their proliferation and capacity to form myotubes.

Previous work also has shown adverse effects of hypoxia on the myogenic response in both adults and fetuses in vivo. In adults, decreased cross-sectional area of myofibers has been shown in humans exposed to chronic (>6 wk) hypobaric oxygen chambers or a high-altitude expedition (41, 42). In both human and sheep fetuses, exposure to hypoxia produces a phenotype of asymmetrical fetal growth or “brain sparing,” whereby the fetus redirects blood to the brain at the expense of splanchnic organs, including the hindlimb and skeletal muscle (30, 43). In sheep models of early onset severe placental insufficiency and IUGR that demonstrated asymmetrical fetal growth, we and others have shown lower femoral arterial blood flow (11, 44), slower hindlimb linear growth rates, smaller muscle weights relative to brain weight and hindlimb weight, and reduced myoblast proliferation rates and myofiber areas (11, 13). However, hypoxemia is only one of many factors, including reduced substrate delivery, that contribute to reduced skeletal muscle growth in the IUGR fetus (11). Brain et al. (45) developed an experimental model of fetal hypoxia by housing pregnant sheep in isobaric hypoxic chambers from 105 to 138 dGA designed to lower maternal and fetal $\text{P}{\text{a}}_{\text{O}}{}_{{}_2}$ by 50% independent of maternal food intake. Fetal hypoxia resulted in asymmetric fetal growth, including lower fetal body weights, shorter hindlimb lengths, and higher brain-to-body weight ratios compared with normoxic fetuses (45). Furthermore, the ratio of oxygen delivery to the ascending relative to the descending aortic circulation was higher, indicating redistribution of oxygen away from the peripheral circulation and toward the brain (46). In the present study, we used serial blood removal to lower the circulating oxygen concentrations in the fetus, which also will include responses to anemia. Despite a relatively shorter period of fetal anemic hypoxemia compared with the above-mentioned models, fetal hindlimb lengths were shorter to a proportional degree as reduced hindlimb muscle weights and myofibers were smaller in the FDS muscle. However, we observed no difference in fetal weight, brain weight, or brain-to-body weight ratio, demonstrating that the hindlimb and the skeletal muscle contained within it are particularly vulnerable to hypoxemia, even after a several day exposure.

Consistent with preserved overall fetal growth, there were no differences in carbon, nitrogen, glucose, or amino acid uptake rates between control and anemic fetuses. This was despite lower oxygen consumption rates in anemic hypoxemic fetuses. Furthermore, fetal protein synthesis and accretion rates were not different between groups. However, protein kinetic rates were measured across the fetus as a whole and do not rule out that particular tissue beds and organs could have higher or lower growth rates relative to others. For example, we previously reported that liver and cardiac weights were higher in anemic hypoxemic fetuses (17, 18), whereas herein we report shorter hindlimbs and lighter hindlimb muscles. In the IUGR fetus as well, we have shown that that hindlimb and muscle growth rates slow as a possible mechanism of sparing substrates for vital organs (11). Additionally, particular muscles were more vulnerable than others within the hindlimb. The BF was most affected in terms of reduced myoblast proliferation, and the FDS was the only muscle with a significant reduction in myofiber size. Although the TA weighed less, it was protected from the effects of anemic hypoxemia on myoblast proliferation and myofiber hypertrophy. Interestingly, the TA was similarly protected from a reduction in myonuclear number in IUGR fetal sheep induced by chronic placental insufficiency (11). Each muscle within the hindlimb has different functional roles, fiber type compositions, and capacity for increased perfusion in response to exercise in adult sheep (23, 24, 47). For example, the FDS is responsible for extending the tarsal joint and flexing the toe joints and ultimately has more than half of its fiber composition as type I oxidative, indicating its role in postural maintenance for grazing (23). The TA is responsible for flexing the tarsal joint and also has a high percentage of type I oxidative fibers in adult sheep muscle (23). The BF is primarily used for ambulation and therefore has a higher percentage of glycolytic fibers (24). The reasons for differential responses among muscles in the hindlimb in response to adverse intrauterine conditions are currently unknown but could be species specific based on the functional importance of that particular muscle in adult life.

Serial blood removal in anemic hypoxemic fetuses also resulted in higher fetal arterial cortisol and norephinephrine concentrations, lower concentrations of select amino acids, and lower IGF-1 concentrations, which may have contributed to slower muscle growth. Elevated catecholamines have been shown to increase muscle protein breakdown in adults (48). However, at least across the whole anemic fetus, we did not observe an increase in protein breakdown rates. This finding is consistent with results from a fetal norepinephrine infusion; fetal protein accretions rates were lower but not as a result of increased protein breakdown (49). Though we did not measure muscle-specific protein kinetics specifically, mRNA expression of regulators of autophagy and protein ubiquitination in skeletal muscle biopsies from anemic hypoxemic fetuses were either similar or reduced compared with control fetuses, providing more evidence that an increase in muscle protein breakdown was not the primary etiology for lower muscle mass and myofiber hypertrophy. Interestingly, a decline in fetal blood O₂ content stimulates an increase in norepinephrine, which in turn suppresses insulin concentrations to slow fetal growth rates, resulting in finely tuned homeostatic control of fetal oxygenation (50–52). However, in this model, insulin concentrations were not different between control and anemic fetuses. The reason for this interaction between norepinephrine and insulin (or lack thereof) is not clear. But, taken together, we find little evidence that elevated norepinephrine contributed significantly to reduced muscle growth.

Lower circulating IGF-1, however, may have had considerable contribution to reduced myofiber hypertrophy and myoblast proliferation. We have shown a strong correlation between hindlimb weight and circulating plasma IGF-1 concentrations in normally growing control and IUGR fetal sheep (11). IGF-1 is a well-known regulator of myogenesis that stimulates both proliferation and hypertrophy (53, 54). In fact, oxygen has been shown to play a critical role in specifying the cellular responses to IGFs (55). It is possible that concurrent reductions in IGF-1 and oxygen availability further contributed to reduced muscle and hindlimb growth in anemic hypoxemic fetuses. However, we did not find differences in the signaling proteins Akt or 4E-BP1 within the IGF-1 signal transduction pathway that are known to stimulate muscle cell growth (56, 57), nor did we find any differences in activated AMPK, which is a sensor of cellular energy status and regulator of skeletal muscle protein synthesis (58). Serial blood withdrawal to produce anemic hypoxemia also resulted in lower circulating concentrations of certain amino acids, which also may have contributed to slower muscle growth as has been shown in adult pigs subjected to hemodialysis to lower circulating amino acids (59). Further investigation into the mechanistic regulation for how anemic hypoxemia reduces fetal muscle growth, either through the downregulation of myogenic regulatory factors, suppression of signaling within alternate anabolic pathways, or from a reduction of substrate availability is warranted, given the negative effects on muscle growth observed in this study and the common occurrence of fetal hypoxemia during human pregnancy (15).

Perspectives and Significance

Ischemic placental disease is a common complication in human pregnancy and includes conditions such as placental insufficiency-induced IUGR, preeclampsia, pregnancy at high altitude, chronic placental abruption, and umbilical cord occlusion that result in fetal hypoxemia and subsequent fetal growth restriction (15, 60, 61). Pregnancy conditions that result in chronic fetal anemia, including maternal alloimmunization, twin anemia polycythemia sequence, placental/fetal tumors, or fetomaternal hemorrhage, will also reduce fetal oxygen content (16). We have shown in an experimental sheep model that fetal anemic hypoxemia for 9 days during late gestation results in muscle type-specific reductions in mass, myofiber size, reduced myoblast proliferation, and downregulation of the myogenic program. Our results are consistent with previous work demonstrating the negative effect of hypoxemia on fetal hindlimb growth (17, 45) and demonstrate that skeletal muscle is more vulnerable to hypoxemia than other fetal organs, even after a limited duration of hypoxemia relative to early human IUGR with onset < 32 weeks gestation (62, 63). This effect on fetal skeletal muscle may carry consequences throughout the life course because the potential for postnatal muscle catch-up growth is limited leading to persistent reductions in muscle mass (5, 7, 64, 65). Our work identifies oxygen availability as a major factor in the regulation of fetal skeletal muscle growth during late gestation.

GRANTS

This work was supported by the National Institute of Health (NIH) Grants R01DK108910 (to S.R.W.), R01HD071068 (to S.S.J.), R01HL142483 (to S.S.J.), R01HD079404 (to L.D.B.), R01DK088139 (to P.J.R.), R01HD093701 (to P.J.R.), S10OD023553 (to L.D.B.), University of Colorado Anschutz Medical Campus Diabetes Research Center Core Services funded by NIH P30-DK116073, and the Ludeman Family Center for Women’s Health Research.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

P.J.R., S.R.W., S.S.J., and L.D.B. conceived and designed research; P.J.R. and L.D.B. performed experiments; P.J.R. and L.D.B. analyzed data; P.J.R., S.R.W., S.S.J., and L.D.B. interpreted results of experiments; P.J.R. and L.D.B. prepared figures; P.J.R. and L.D.B. drafted manuscript; P.J.R., S.R.W., S.S.J., and L.D.B. edited and revised manuscript; P.J.R., S.R.W., S.S.J., and L.D.B. approved final version of manuscript.