Hypoxic stress in diabetic pregnancy contributes to impaired embryo gene expression and defective development by inducing oxidative stress.

We have shown that neural tube defects (NTD) in a mouse model of diabetic embryopathy are associated with deficient expression of Pax3, a gene required for neural tube closure. Hyperglycemia-induced oxidative stress is responsible. Before organogenesis, the avascular embryo is physiologically hypoxic (2-5% O(2)). Here we hypothesized that, because O(2) delivery is limited at this stage of development, excess glucose metabolism could accelerate the rate of O(2) consumption, thereby exacerbating the hypoxic state. Because hypoxia can increase mitochondrial superoxide production, excessive hypoxia may contribute to oxidative stress. To test this, we assayed O(2) flux, an indicator of O(2) availability, in embryos of glucose-injected hyperglycemic or saline-injected mice. O(2) flux was reduced by 30% in embryos of hyperglycemic mice. To test whether hypoxia replicates, and hyperoxia suppresses, the effects of maternal hyperglycemia, pregnant mice were housed in controlled O(2) chambers on embryonic day 7.5. Housing pregnant mice in 12% O(2), or induction of maternal hyperglycemia (>250 mg/dl), decreased Pax3 expression fivefold, and increased NTD eightfold. Conversely, housing pregnant diabetic mice in 30% O(2) significantly suppressed the effect of maternal diabetes to increase NTD. These effects of hypoxia appear to be the result of increased production of mitochondrial superoxide, as indicated by assay of lipid peroxidation, reduced glutathione, and H(2)O(2). Further support of this interpretation was the effect of antioxidants, which blocked the effects of maternal hypoxia, as well as hyperglycemia, on Pax3 expression and NTD. These observations suggest that maternal hyperglycemia depletes O(2) in the embryo and that this contributes to oxidative stress and the adverse effects of maternal hyperglycemia on embryo development.

Life at acidic pH imposes an increased energetic cost for a eukaryotic acidophile.

Organisms growing in acidic environments, pH<3, would be expected to possess fundamentally different molecular structures and physiological controls in comparison with similar species restricted to neutral pH. We begin to investigate this premise by determining the magnitude of the transmembrane electrochemical H+ gradient in an acidophilic Chlamydomonas sp. (ATCC PRA-125) isolated from the Rio Tinto, a heavy metal laden, acidic river (pH 1.7-2.5). This acidophile grows most rapidly at pH 2 but is capable of growth over a wide pH range (1.5-7.0), while Chlamydomonas reinhardtii is restricted to growth at pH>or=3 with optimal growth between pH 5.5 and 8.5. With the fluorescent H+ indicator, 2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF), we show that the acidophilic Chlamydomonas maintains an average cytosolic pH of 6.6 in culture medium at both pH 2 and pH 7 while Chlamydomonas reinhardtii maintains an average cytosolic pH of 7.1 in pH 7 culture medium. The transmembrane electric potential difference of Chlamydomonas sp., measured using intracellular electrodes at both pH 2 and 7, is close to 0 mV, a rare value for plants, animals and protists. The 40,000-fold difference in [H+] could be the result of either active or passive mechanisms. Evidence for active maintenance was detected by monitoring the rate of ATP consumption. At the peak, cells consume about 7% more ATP per second in medium at pH 2 than at pH 7. This increased rate of consumption is sufficient to account for removal of H+ entering the cytosol across a membrane with relatively high permeability to H+ (7x10(-8) cm s-1). Our results indicate that the small increase in the rate of ATP consumption can account for maintenance of the transmembrane H+ gradient without the imposition of cell surface H+ barriers.

Direct measurement of glucose gradients and mass transport within islets of Langerhans.

A glucose oxidase-based glucose microsensor (<10 micro m tip diameter) was used to measure the glucose concentration within single islets under static conditions and during step changes in glucose level. The sensors had response times of 4.1+/-0.5s (n=7) and sensitivities of 8.7+/-1.8 pA/mM (n=11). The sensors performed independent of oxygen up to 15 mM glucose as long as the oxygen level was > 70 mm Hg. Spatially resolved glucose measurements revealed a glucose gradient around and inside single islets. From measurement of the glucose gradient, a glucose consumption rate of 0.48+/-0.14 pmol/nL islet/min (n=6) and an intraislet glucose diffusion coefficient of 3.8 x 10(-7)cm(2)/s were determined. The measurement of the gradient demonstrates that not all cells within an islet in culture are exposed to the same glucose concentration. The sensor was also used to measure the time required for intraislet glucose concentration to reach steady state following a step increase in glucose concentration from 3 to 10 mM at the islet surface. At a depth of 70 micro m inside an islet, glucose reached steady state in 180+/-7s (n=7) for islets with a diameter of 180-220 micro m (smaller islets reach steady-state faster). In the presence of 10 mM mannoheptulose, an inhibitor of glucokinase, the equilibration time was reduced to 122+/-11s (n=6), indicating that glucose utilization by glycolysis limited the time required for glucose to diffuse into the islets. The long times to reach steady state and presence of glucose gradients are important in interpreting data from experiments involving islets in culture.

Metabolic oscillations in beta-cells.

Whereas the mechanisms underlying oscillatory insulin secretion remain unknown, several models have been advanced to explain if they involve generation of metabolic oscillations in beta-cells. Evidence, including measurements of oxygen consumption, glucose consumption, NADH, and ATP/ADP ratio, has accumulated to support the hypothesis that energy metabolism in beta-cells can oscillate. Where simultaneous measurements have been made, these oscillations are well correlated with oscillations in intracellular [Ca(2+)] and insulin secretion. Considerable evidence has been accumulated to suggest that entry of Ca(2+) into cells can modulate metabolism both positively and negatively. The main positive effect of Ca(2+) is an increase in oxygen consumption, believed to involve activation of mitochondrial dehydrogenases. Negative feedback by Ca(2+) includes decreases in glucose consumption and decreases in the mitochondrial membrane potential. Ca(2+) also provides negative feedback by increasing consumption of ATP. The negative feedback provided by Ca(2+) provides a mechanism for generating oscillations based on a model in which glucose stimulates a rise in ATP/ADP ratio that closes ATP-sensitive K(+) (K(ATP)) channels, thus depolarizing the cell membrane and allowing Ca(2+) entry through voltage-sensitive channels. Ca(2+) entry reduces the ATP/ADP ratio and allows reopening of the K(ATP) channel.