Information

14: Chemolithotrophy & Nitrogen Metabolism - Biology

14: Chemolithotrophy & Nitrogen Metabolism - Biology



We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

Chemolithotrophy

Chemolithotrophy is the oxidation of inorganic chemicals for the generation of energy. The process can use oxidative phosphorylation, just like aerobic and anaerobic respiration, but now the substance being oxidized (the electron donor) is an inorganic compound. The electrons are passed off to carriers within the electron transport chain, generating a proton motive force that is used to generate ATP with the help of ATP synthase.

Chemolithotrophy Pathways.

Electrons donors

Chemolithotrophs use a variety of inorganic compounds as electron donors, with the most common substances being hydrogen gas, sulfur compounds (such as sulfide and sulfur), nitrogen compounds (such as ammonium and nitrite), and ferrous iron.

  • Hydrogen oxidizers – these organisms oxidize hydrogen gas (H2) with the use of a hydrogenase enzyme. Both aerobic and anaerobic hydrogen oxidizers exist, with the aerobic organisms eventually reducing oxygen to water.
  • Sulfur oxidizers – as a group these organisms are capable of oxidizing a wide variety of reduced and partially reduced sulfur compounds such as hydrogen sulfide (H2S), elemental sulfur (S0), thiosulfate (S2O32-), and sulfite (SO32-). Sulfate (SO42-) is frequently a by-product of the oxidation. Often the oxidation occurs in a stepwise fashion with the help of the sulfite oxidase enzyme.
  • Nitrogen oxidizers – the oxidation of ammonia (NH3) is performed as a two-step process by nitrifying microbes, where one group oxidizes ammonia to nitrite (NO2-) and the second group oxidizes the nitrite to nitrate (NO3-). The entire process is known as nitrification and is performed by small groups of aerobic bacteria and archaea, often found living together in soil or in water systems.
  • Iron oxidizers – these organisms oxidize ferrous iron (Fe2+) to ferric iron (Fe3+). Since Fe2+ has such a positive standard reduction potential, the bioenergetics are not extremely favorable, even using oxygen as a final electron acceptor. The situation is made more difficult for these organisms by the fact that Fe2+ spontaneously oxidizes to Fe3+ in the presence of oxygen; the organisms must use it for their own purposes before that happens.

Electron acceptors

Chemolithotrophy can occur aerobically or anaerobically. Just as with either type of respiration, the best electron acceptor is oxygen, to create the biggest distance between the electron donor and the electron acceptor. Using a non-oxygen acceptor allows chemolithotrophs to have greater diversity and the ability to live in a wider variety of environments, although they sacrifice energy production.

Amount of ATP generated

Just as both the electron donors and acceptors can vary widely for this group of organisms, the amount of ATP generated for their efforts will vary widely as well. They will not make as much ATP as an organism using aerobic respiration, since the largest ΔE0’ is found using glucose as an electron donor and oxygen as an electron acceptor. But how much less than 32 molecules of ATP greatly depends upon the actual donor and acceptor being used. The smaller the distance between the two, the less ATP that will be formed.

Chemolithoautotrophs vs chemolithoheterotrophs

Most chemolithotrophs are autotrophs (chemolithoautotrophs), where they fix atmospheric carbon dioxide to assemble the organic compounds that they need. These organisms require both ATP and reducing power (i.e. NADH/NADPH) in order to ultimately convert the oxidized molecule CO2 into a greatly reduced organic compound, like glucose. If a chemolithoautotroph is using an electron donor with a higher redox potential than NAD+/NADP, they must use reverse electron flow to push electrons back up the electron tower. This is energetically unfavorable to the cell, consuming energy from the proton motive force to drive electrons in a reverse direction back through the ETC.

Some microbes are chemolithoheterotrophs, using an inorganic chemical for their energy and electron needs, but relying on organic chemicals in the environment for their carbon needs. These organisms are also called mixotrophs, since they require both inorganic and chemical compounds for their growth and reproduction.

Nitrogen Metabolism

The nitrogen cycle depicts the different ways in which nitrogen, an essential element for life, is used and converted by organisms for various purposes. Much of the chemical conversions are performed by microbes as part of their metabolism, performing a valuable service in the process for other organisms in providing them with an alternate chemical form of the element.

Nitrogen Cycle.

Nitrogen Fixation

Nitrogen fixation describes the conversion of the relatively inert dinitrogen gas (N2) into ammonia (NH3), a much more useable form of nitrogen for most life forms. The process is performed by diazotrophs, a limited number of bacteria and archaea that can grow without an external source of fixed nitrogen, because of their abilities. Nitrogen fixation is an essential process for Earth’s organisms, since nitrogen is a required component of various organic molecules, such as amino acids and nucleotides. Plants, animals, and other organisms rely on bacteria and archaea to provide nitrogen in a fixed form, since no eukaryote is known that can fix nitrogen.

Nitrogen fixation is an extremely energy and electron intensive process, in order to break the triple bond in N2 and reduce it to NH3. It requires a particular enzyme known as nitrogenase, which is inactivated by O2. Thus, nitrogen fixation must take place in an anaerobic environment. Aerobic nitrogen-fixing organisms must devise special conditions or arrangements in order to protect their enzyme. Nitrogen-fixing organisms can either exist independently or pair up with a plant host:

  1. Symbiotic nitrogen-fixing organisms: these bacteria partner up with a plant, to provide them with an environment appropriate for the functioning of their nitrogenase enzyme. The bacteria live in the plant’s tissue, often in root nodules, fixing nitrogen and sharing the results. The plant provides both the location to fix nitrogen, as well as additional nutrients to support the energy-taxing process of nitrogen fixation. It has been shown that the bacteria and the host exchange chemical recognition signals that facilitate the relationship. One of the best known bacteria in this category is Rhizobium, which partners up with plants of the legume family (clover, soybeans, alfalfa, etc).
  2. Free-living nitrogen-fixing organisms: these organisms, both bacteria and archaea, fix nitrogen for their own use that ends up being shared when the organisms dies or is ingested. Free-living nitrogen-fixing organisms that grow anaerobically do not have to worry about special adaptations for their nitrogenase enzyme. Aerobic organisms must make adaptations. Cyanobacteria, a multicellular bacterium, make specialized cells known as heterocystsin which nitrogen fixation occurs. Since Cyanobacteria produce oxygen as part of their photosynthesis, an anoxygenic version occurs within the heterocyst, allowing the nitrogenase to remain active. The heterocysts share the fixed nitrogen with surrounding cells, while the surrounding cells provide additional nutrients to the heterocysts.

Assimilation

Assimilation is a reductive process by which an inorganic form of nitrogen is reduced to organic nitrogen compounds such as amino acids and nucleotides, allowing for cellular growth and reproduction. Only the amount needed by the cell is reduced. Ammonia assimilation occurs when the ammonia (NH3)/ammonium ion (NH4+) formed during nitrogen fixation is incorporated into cellular nitrogen. Assimilative nitrate reduction is a reduction of nitrate to cellular nitrogen, in a multi-step process where nitrate is reduced to nitrite then ammonia and finally into organic nitrogen.

Nitrification

As mentioned above, nitrification is performed by chemolithotrophs using a reduced or partially reduced form of nitrogen as an electron donor to obtain energy. ATP is gained by the process of oxidative phosphorylation, using a ETC, PMF, and ATP synthase.

Denitrification

Denitrification refers to the reduction of NO3- to gaseous nitrogen compounds, such as N2. Denitrifying microbes perform anaerobic respiration, using NO3- as an alternate final electron acceptor to O2. This is a type of dissimilatory nitrate reduction where the nitrate is being reduced during energy conservation, not for the purposes of making organic compounds. This produces large amounts of excess byproducts, resulting in the loss of nitrogen from the local environment to the atmosphere.

Anammox

Anammox or anaerobic ammonia oxidation is performed by marine bacteria, relatively recently discovered, that utilize nitrogen compounds as both electron acceptor and electron donor. Ammonia is oxidized anaerobically as the electron donor while nitrite is utilized as the electron acceptor, with dinitrogen gas produced as a byproduct. The reactions occur within the anammoxosome, a specialized cytoplasmic structure which constitutes 50-70% of the total cell volume. Just like denitrification, the anammox reaction removes fixed nitrogen from a local environment, releasing it to the atmosphere.

Key Words

chemolithotrophy, hydrogen oxidizers, hydrogenase, sulfur oxidizers, sulfite oxidase, nitrogen oxidizers, nitrification, iron oxidizers, chemolithoautotroph, reverse electron flow, chemolithoheterotroph, mixotroph, nitrogen fixation, diazotroph, nitrogenase, symbiotic nitrogen-fixing organisms, Rhizobium, legume, free-living nitrogen-fixing organisms, Cyanobacteria, heterocyst, assimilation, ammonia assimilation, assimilative nitrate reduction, denitrification, dissimilatory nitrate reduction, anammox, anaerobic ammonia oxidation, anammoxosome.

Study Questions

  1. What is chemolithotrophy?
  2. What are the most common electron donors and acceptors for chemolithotrophs? How does their amount of ATP produced compare to chemoorganotrophs?
  3. How do chemolithoautotrophs and chemolithoheterotrophs differ? What is the reverse electron flow and how/why is it used by some chemolithoautotrophs?
  4. What roles do bacteria/archaea play in the nitrogen cycle? How are different nitrogen compounds used in their metabolism?
  5. What is required for nitrogen fixation? How do free living nitrogen fixers and plant associated nitrogen fixers differ? How do Rhizobium and Cyanobacteria protect their nitrogenase from oxygen?
  6. What are the different mechanisms of nitrogen metabolism? What conversion is occurring for each? What is the purpose of each and how does it relate to the organism’s metabolism?

Nitrogen Cycle.

Nitrogen Fixation

Nitrogen fixation describes the conversion of the relatively inert dinitrogen gas (N2) into ammonia (NH3), a much more useable form of nitrogen for most life forms. The heterocysts share the fixed nitrogen with surrounding cells, while the surrounding cells provide additional nutrients to the heterocysts.

Assimilation

Assimilation is a reductive process by which an inorganic form of nitrogen is reduced to organic nitrogen compounds such as amino acids and nucleotides, allowing for cellular growth and reproduction. Assimilative nitrate reduction is a reduction of nitrate to cellular nitrogen, in a multi-step process where nitrate is reduced to nitrite then ammonia and finally into organic nitrogen.

Nitrification

As mentioned above, nitrification is performed by chemolithotrophs using a reduced or partially reduced form of nitrogen as an electron donor to obtain energy. ATP is gained by the process of oxidative phosphorylation, using a ETC, PMF, and ATP synthase.

Denitrification

Denitrification refers to the reduction of NO3- to gaseous nitrogen compounds, such as N2. This produces large amounts of excess byproducts, resulting in the loss of nitrogen from the local environment to the atmosphere.

Anammox

Anammox or anaerobic ammonia oxidation is performed by marine bacteria, relatively recently discovered, that utilize nitrogen compounds as both electron acceptor and electron donor. Just like denitrification, the anammox reaction removes fixed nitrogen from a local environment, releasing it to the atmosphere.

Key Words

chemolithotrophy, hydrogen oxidizers, hydrogenase, sulfur oxidizers, sulfite oxidase, nitrogen oxidizers, nitrification, iron oxidizers, chemolithoautotroph, reverse electron flow, chemolithoheterotroph, mixotroph, nitrogen fixation, diazotroph, nitrogenase, symbiotic nitrogen-fixing organisms, Rhizobium, legume, free-living nitrogen-fixing organisms, Cyanobacteria, heterocyst, assimilation, ammonia assimilation, assimilative nitrate reduction, denitrification, dissimilatory nitrate reduction, anammox, anaerobic ammonia oxidation, anammoxosome.

Study Questions

  1. What is chemolithotrophy?
  2. What are the most common electron donors and acceptors for chemolithotrophs? How does their amount of ATP produced compare to chemoorganotrophs?
  3. How do chemolithoautotrophs and chemolithoheterotrophs differ? What is the reverse electron flow and how/why is it used by some chemolithoautotrophs?
  4. What roles do bacteria/archaea play in the nitrogen cycle? How are different nitrogen compounds used in their metabolism?
  5. What is required for nitrogen fixation? How do free living nitrogen fixers and plant associated nitrogen fixers differ? How do Rhizobium and Cyanobacteria protect their nitrogenase from oxygen?
  6. What are the different mechanisms of nitrogen metabolism? What conversion is occurring for each? What is the purpose of each and how does it relate to the organism’s metabolism?

Coordination of bacterial proteome with metabolism by cyclic AMP signalling

The cyclic AMP (cAMP)-dependent catabolite repression effect in Escherichia coli is among the most intensely studied regulatory processes in biology. However, the physiological function(s) of cAMP signalling and its molecular triggers remain elusive. Here we use a quantitative physiological approach to show that cAMP signalling tightly coordinates the expression of catabolic proteins with biosynthetic and ribosomal proteins, in accordance with the cellular metabolic needs during exponential growth. The expression of carbon catabolic genes increased linearly with decreasing growth rates upon limitation of carbon influx, but decreased linearly with decreasing growth rate upon limitation of nitrogen or sulphur influx. In contrast, the expression of biosynthetic genes showed the opposite linear growth-rate dependence as the catabolic genes. A coarse-grained mathematical model provides a quantitative framework for understanding and predicting gene expression responses to catabolic and anabolic limitations. A scheme of integral feedback control featuring the inhibition of cAMP signalling by metabolic precursors is proposed and validated. These results reveal a key physiological role of cAMP-dependent catabolite repression: to ensure that proteomic resources are spent on distinct metabolic sectors as needed in different nutrient environments. Our findings underscore the power of quantitative physiology in unravelling the underlying functions of complex molecular signalling networks.


Ammonia assimilation and reassimilation

Glutamine synthetase (GS EC 6.3.1.2) was first purified and characterized from plants in 1956. One particular important characteristic is its high affinity for ammonia and thus its ability to incorporate ammonia efficiently into organic combination. Originally, glutamine was considered to donate its amide N only into a limited number of compounds. However, the discovery of NAD(P)H glutamate synthase in bacteria ( Tempest et al., 1970) and later ferredoxin‐dependent glutamate synthase in plants ( Lea and Miflin, 1974) established a route, the glutamate synthase cycle, for NH3 2 to enter into organic compounds via its assimilation by GS. Evidence based on labelling kinetics, use of inhibitors, in organello studies, and genetics established that this was the major route of primary nitrogen assimilation in plants ( Miflin and Lea, 1980).

During the growth and development of plants, nitrogen is moved into and out of proteins in the different organs and transported between organs in a limited number of transport compounds. Some of the organic nitrogen is moved between compounds via the activity of transaminases and glutamine‐amide transferases, but a significant portion is released as NH3 and reassimilated via GS. For example, asparagine is a significant component of seed storage proteins in legumes and a major transport compound in cereals. It is metabolized to ammonia and aspartate via the action of asparaginase. Similarly ureides, such as allantoin, play an important role in N transport in legumes and their organic N is released as NH3 via the action of urease. Thus, over the life of a plant, nitrogen is released as NH3 and refixed several times (for a detailed description of these processes see Lea and Miflin, 1980). This flux through NH3 and GS is dwarfed in C3 plants by the flux of NH3 released by glycine decarboxylase during photorespiration. This could be an order of magnitude more than the rate of primary assimilation. Biochemical and genetic experiments have shown that this NH3 is also refixed via GS ( Keys et al., 1978 Somerville and Ogren, 1982 Wallsgrove et al., 1987).

Overall GS acts at the centre of nitrogen flow as depicted by the scheme in the centre of Fig. 1 . This central position of GS raises a number of crucial questions: How does the plant maintain C/N balance? How is GS distributed? How is GS activity regulated? Does glutamine regulate metabolism? Does GS regulate development? Is our knowledge of GS sufficient to be useful? Can we improve plants agronomically by modifying GS?

The central role of GS in the complex matrix of plant N metabolism. The central scheme encompasses the total role of GS. The boxes around the outside indicate the matrix of various locations and environments in which GS may be operating. The direction of the flow of N (and thus the arrows) will depend on which part of the matrix is under consideration. Thus in the developing seed the flux will be from incoming transport compounds towards proteins whilst in the germinating seed the flow will be in the reverse direction.

The central role of GS in the complex matrix of plant N metabolism. The central scheme encompasses the total role of GS. The boxes around the outside indicate the matrix of various locations and environments in which GS may be operating. The direction of the flow of N (and thus the arrows) will depend on which part of the matrix is under consideration. Thus in the developing seed the flux will be from incoming transport compounds towards proteins whilst in the germinating seed the flow will be in the reverse direction.


Interdependence of CO2 and inorganic nitrogen on crassulacean acid metabolism and efficiency of nitrogen use by Littorella uniflora (L.) Aschers

The hypothesis is tested that crassulacean acid metabolism (CAM) in isoetids is a mechanism which not only conserves inorganic carbon but also plays a role in nitrogen economy of the plants. This hypothesis was tested in an outdoor experiment, where Littorella uniflora (L.) Aschers. were grown at two CO2 and five inorganic nitrogen concentrations in a crossed factorial design. The growth of Littorella responded positively to enhanced nitrogen availability at high but not at low CO2 indicating that growth was limited by nitrogen at high CO2 only. For the nitrogen-limited plants, the capacity for CAM (CAMcap) increased with the degree of nitrogen limitation of growth and an inverse coupling between CAM and tissue-N was found. Although this might indicate a role of CAM in economizing on nitrogen in Littorella, the hypothesis was rejected for the following reasons: (1) although CAMcap was related to tissue-N no relationship between tissue-N and ambient CAM activity (CAMambient) was found whereas a close relationship would be expected if CAM was regulated by nitrogen availability (2) the photosynthetic nitrogen use efficiency for high CO2-grown plants declined with increased CAMambient and with CAMcap and (3) growth per unit tissue-N per unit time declined with increased CAMambient and CAMcap.


Exchange of Gases at the Alveolar Surface

Gas exchange between air and blood occurs at the alveoli. Gaseous exchange occurs by the process of simple diffusion between the alveolar air and the deoxygenated blood in capillaries. Due to the existing pressure difference of oxygen and carbon dioxide between the alveoli and the blood capillaries, oxygen diffuses from alveolar air to the capillary blood, whereas carbon dioxide diffuses from capillary blood to the alveolar air. Oxygenated blood is taken from the lungs to the heart by pulmonary vein.

Volume exchanged during breathing:

Table Showing Air Volume Exchanged During Breathing
Tidal volume (TV)Volume of air inhaled and exhaled without any noticeable effort (normal breathing)500mL
Vital capacity (VC)Volume of air that can be maximally breathed out after a maximum inspiration (VC = IRV + TV + ERV)3400 – 4800 mL
Inspiratory reserve volume (IRV)Volume of air that can be taken in by forced inspiration over and above the normal inspiration2000 – 3000 mL
Expiratory reserve volume (ERV)Volume of air that can be expelled by forced expiration over and above the normal expiration.1000 mL
Residual volume (RV)Volume of air that control be forced out even on forced expiration. This is the air that remains in the lungs and in the air passage.1000 – 1500 mL
Total lung capacitySum of all lung volumes (maximum air that remains in the lungs after a maximum inhalation)5500 – 6000 mL

Vital capacity may be highly reduced in smokers and people suffering from tuberculosis. Athletes and singers on the other hand have higher vital capacity.


Results

Activities of the ornithine-urea cycle enzymes and glutamine synthetase from the liver

No carbamoyl phosphate synthetase I or III activity was detected (detection limit=0.001 μmol min -1 g -1 tissue N=4) in the liver tissue from A. testudineus immersed in water. As for ornithine transcarbamoylase, argininosuccinate synthetase + lyase, argainase and glutamine synthetase, only very low activities (0.10±0.02,0.006±0.001, 3.3±0.9 and 0.054±0.012 μmol min -1 g -1 tissue, respectively N=4) were detected.

Effects of emersion on rates of ammonia and urea excretion

With daily change of water, the ammonia concentration in the thin film of water (80 ml) reached approximately 4-5 mmol l -1 after 24 h throughout the 4-day period of emersion. This indicates that the rate of ammonia excretion in A. testudineus during emersion was high. Indeed,the daily ammonia excretion rates (N=12) in A. testudineuson days 1, 3 and 4 of emersion were not significantly different from those of the control immersed in water (Fig. 1A). Surprisingly, on day 2, the daily ammonia excretion rate of the fish exposed to terrestrial conditions was significantly greater than that of the control. When summed together over a 2-day period, however, the excretion rate of 11.5±2.1 μmol 2 days -1 g -1 was not significantly different from the control value of 13.9± 0.7μmol 2 days -1 g -1 . Similarly, the total rate of ammonia excretion in the experimental (24.1±4.6 μmol 4 days -1 g -1 ) and control (32.2±1.9 μmol 4 days -1 g -1 ) fish over a 4-day period were comparable. Upon re-immersion on day 5, the daily ammonia excretion rate (N=6) of the experimental fish was not significantly different from that of the immersed control (Fig. 1A).

By contrast, in spite of significant increases, the urea concentration in the thin film of water remained relatively low throughout the 4-day emersion period. Emersion led to a significant decrease in the daily urea excretion rate on day 1, but had no significant effect on urea excretion thereafter(Fig. 1B). There was no significant change in urea excretion in fish during re-immersion on day 5 as compared with day 1 or day 5 controls (Fig. 1B).

Rates (μmol day -1 g -1 fish) of (A) ammonia and (B)urea excretion in Anabas testudineus immersed in water (control C)for 5 days (N=12 for days 1 to 4 and N=6 for day 5) or exposed to terrestrial conditions (T) for 4 days (N=12) followed by 1 day (N=6) of re-immersion in water. Values are means ± s.e.m. * Significantly different from the corresponding freshwater control condition. † Significantly different from the corresponding day 1, terrestrial condition.

Rates (μmol day -1 g -1 fish) of (A) ammonia and (B)urea excretion in Anabas testudineus immersed in water (control C)for 5 days (N=12 for days 1 to 4 and N=6 for day 5) or exposed to terrestrial conditions (T) for 4 days (N=12) followed by 1 day (N=6) of re-immersion in water. Values are means ± s.e.m. * Significantly different from the corresponding freshwater control condition. † Significantly different from the corresponding day 1, terrestrial condition.

In a separate set of experiments in which daily change of water was omitted for a 2-day period, the ammonia concentration (N=5) in the thin film of water reached 6.68±1.11 and 13.2±2.1 mmol l -1 at the end of day 1 and day 2, respectively. The respective daily ammonia excretion rates were 9.98±1.92 and 8.64±1.85 μmol day -1 g -1 fish, which were not significantly different from the corresponding values (7.45±1.32 and 7.54±1.21 μmol day -1 g -1 fish, respectively) of the control fish(N=5) immersed in water.

Ammonia concentrations in water samples collected from branchial or cutaneous surfaces of fish after 15 min or 24 h of emersion

The ammonia concentration in water samples collected from the branchial surface of the air-facing side of the fish after 24 h of emersion(N=5) was not significantly different from those of the control exposed to terrestrial conditions for 15 min during anaesthesia (N=5 Fig. 2). However, for the water-facing side of the experimental fish, the ammonia concentration in the branchial water increased to 21.5±2.4 mmol l -1 , which was significantly greater than that of the control fish(Fig. 2). In water samples collected from the air-facing cutaneous surface of the experimental fish, the ammonia concentration varied greatly and was not significantly different from the control value (Fig. 2). By contrast, the ammonia concentration (20.8±3.5 mmol l -1 ) in water samples collected from the waterfacing cutaneous surface was significantly greater than that of the control. The concentration of ammonia in the thin film of water at the bottom (80 ml) was 5.32±0.87 mmol l -1 .

Effects of environmental ammonia (12 mmol l -1 ) on rates of ammonia and urea excretion

The ammonia excretion rate of A. testudineus (N=7) in normal freshwater without NH4Cl was 7.12±1.04 μmol day -1 g -1 fish. At the beginning of the experiment, the ambient ammonia concentration without any fish was 12.3 mmol l -1 . With an initial pH of 7.0, the NH4 + and NH3concentrations were calculated to be 12.19 and 0.11 mmol l -1 ,respectively. In comparison, the concentrations of NH4 + and NH3 in the plasma were 0.188 and 0.012 mmol l -1 ,respectively, taking the plasma ammonia concentration to be 0.2 mmol l -1 (from Table 1)and the blood pH to be 7.6 (Y.K.I., unpublished results). So, both the NH4 + and NH3 gradients were directed inwardsbut, surprisingly, the experimental fish could excrete ammonia against such a large ammonia gradient. As a result, the concentration of ammonia in the external medium increased to 13.1±0.2 mmol l -1 at the end of day 1. A simple calculation reveals that the ammonia excretion rate decreased significantly to 4.31±0.81 μmol day -1 g -1 fish(N=7) during this 24 h period. On day 2, the ambient ammonia concentration increased further to 14.7±0.3 mmol l -1 , and the ammonia excretion rate (8.32±1.44 μmol day -1 g -1 fish N=7) returned back to a level comparable with the initial control value.

Concentrations (mmol l -1 ) of ammonia in water samples collected from the branchial or cutaneous surfaces of the air-facing side or the water-facing side of Anabas testudineus immersed in water and anaesthetized for 15 min on land (control C) or exposed to terrestrial conditions for 24 h followed with 15 min of anesthetization on land (T). * Significantly different from the corresponding control condition.

Concentrations (mmol l -1 ) of ammonia in water samples collected from the branchial or cutaneous surfaces of the air-facing side or the water-facing side of Anabas testudineus immersed in water and anaesthetized for 15 min on land (control C) or exposed to terrestrial conditions for 24 h followed with 15 min of anesthetization on land (T). * Significantly different from the corresponding control condition.

Effects of emersion on the contents of ammonia, urea and FAAs

Ammonia in the muscle and liver of A. testudineus (N=6)increased significantly during emersion and peaked at 4.06 μmol g -1 on day 2 (4.5-fold of the control value) and 10.7 μmol g -1 on day 1 (5.5-fold of the control value), respectively(Table 1). The ammonia concentration in the plasma of A. testudineus increased significantly during the first 2 days of emersion, but returned to the control level thereafter (Table 1). Although urea also increased significantly in the muscle and liver of A. testudineus during emersion, the peak levels (0.79 μmol g -1 and 0.81 μmol g -1 on day 2, respectively) attained were much lower than those of ammonia (Table 2).

Emersion led to significant increases in isoleucine, leucine,phenylalanine, tyrosine and valine in the muscle of A. testudineus(N=4) (Table 3). By contrast, the aspartate and glutamine content of muscle in fish emersed for 2 or 4 days were significantly lower than the corresponding control value. Although emersion had no significant effect on the muscle TFAA, there was a significant increase in the muscle TEFAA in fish exposed to 2 days of emersion. Two days of emersion led to a significant increase (8.8-fold) in the lysine content of the liver of A. testudineus(Table 4). However, 4 days of emersion resulted in significant increases in arginine and phenylalanine in the liver. In addition, there was a significant decrease in the glutamate in the liver of fish after 2 or 4 days of emersion. After 2 days of emersion, the brain glutamine content of A. testudineus increased significantly by 2.5-fold (Table 5). There were also significant decreases in alanine, aspartate and glutamate in the brain of fish during the first 2 days of emersion. However, the TFAA and TEFAA content in the liver and brain of A. testudineus were unaffected by emersion.

Effects of 10 min of forced exercise on land

Forced exercise for 10 min on land led to significant increases in excretion of ammonia (0.103±0.009 μmol 10 min -1 g -1 fish) and urea (0.015±0.002 μmol 10 min -1 g -1 fish) as compared with the control on land but without exercise(0.047±0.014 μmol g -1 fish, and 0.003±0.001μmol g -1 fish, respectively). As a result, the concentration of ammonia in the thin film of water in the container with the exercised fish reached 1.79±0.05 mmol l -1 which was significantly higher than that of the control (0.86±0.20 mmol l -1 ). In addition,there were significant increases in ammonia, alanine, lysine and TEFAA in the muscle of fish after forced exercise on land as compared with the control fish exposed to terrestrial conditions for 10 min without disturbance(Table 6). However, forced exercise on land did not have any significant effect on ATP, ADP, AMP,glucose, glycogen, lactate and succinate in the muscle of A. testudineus (Table 6).

Rate of O2 consumption

For the control fish in freshwater (N=4), the O2consumption rates in water and air were 1.38±0.13 and 3.10± 0.54μmol h -1 g -1 fish, respectively. Taken together, the total O2 consumption rate was 4.47±0.48 μmol h -1 g -1 fish. For fish exposed to terrestrial conditions(N=4 for each group) for 1, 24 or 48 h, the O2 consumption rates in air were 5.25±0.61, 4.67±0.22, and 6.43±0.53μmol h -1 g -1 fish, respectively, and the value obtained from those exposed to terrestrial conditions for 48 h of emersion was significantly greater than that of the immersed control.

Construction of a balance sheet of ammonia and urea-N excretion and ammonia and urea-N retention in a 50 g fish

A 50 g fish contained approximately 30 g of muscle and 1 g of liver. Based on our results, a balance sheet was constructed for changes in nitrogen excretion and changes in ammonia-N and urea-N content in a 50 g fish after 2 or 4 days of immersion or emersion (Table 7). Although there was no significant change in the overall ammonia excretion rate during 4 days of emersion, we took into consideration the small changes involved and presented them in Table 7. After considering the amount of ammonia-N and urea-N stored in the muscle and liver, it becomes apparent that there was an increase in nitrogen production in A. testudineus after 2 or 4 days of emersion.

A balance sheet of changes in ammonia and urea excretion and changes in ammonia and urea content of the muscle and liver of a 50 g Anabas testudineus after 2 or 4 days of immersion (control) or emersion

. Day 2 . . . Day 4 . . .
. Immersion . Emersion . Difference . Immersion . Emersion . Difference .
In a 50 g fish
Ammonia-N excreted 575 697 +122 1205 1608 +403
Urea-N excreted 77 53 −24 142 116 −26
Reduction in nitrogenous excretion (A) +98 +377
In 30 g muscle
Ammonia-N retained 26 122 +96 32 73 +41
Urea-N retained 7 47 +40 3 6 +3
In 1 g liver
Ammonia-N retained 4 9 +5 3 6 +3
Urea-N retained 0.3 1.6 +1.3 0.6 1.4 +0.8
Increase in nitrogenous accumulation (B) +142 +48
(A) + (B) +240 +425
. Day 2 . . . Day 4 . . .
. Immersion . Emersion . Difference . Immersion . Emersion . Difference .
In a 50 g fish
Ammonia-N excreted 575 697 +122 1205 1608 +403
Urea-N excreted 77 53 −24 142 116 −26
Reduction in nitrogenous excretion (A) +98 +377
In 30 g muscle
Ammonia-N retained 26 122 +96 32 73 +41
Urea-N retained 7 47 +40 3 6 +3
In 1 g liver
Ammonia-N retained 4 9 +5 3 6 +3
Urea-N retained 0.3 1.6 +1.3 0.6 1.4 +0.8
Increase in nitrogenous accumulation (B) +142 +48
(A) + (B) +240 +425

Conclusion and perspectives

In the past decade, tremendous research efforts have uncovered key metabolic specificities and Achilles heels of cancer cells, including AML cells. These studies strongly suggest that myeloid leukemias are metabolic disorders and should be regarded in this light for metabolic-based personalized medicine treatments as well as for monitoring clinical responses to treatment. Several studies have further shown that AML cells, like other normal and cancer cells, are able to undergo compensatory metabolic and energetic adaptations in response to the inhibition of metabolic pathways, indicating that AML cells display complex metabolic capacities and flexibility that limit sustained drug efficacy, especially when challenged by chemotherapeutic drugs. However, targeting metabolic flexibility per se is not a feasible approach. By contrast, non-exclusive therapeutic strategies, which impede this metabolic flexibility by targeting its consequence(s), such as mitochondrial dependency, blocking the utilization of nutrients from the microenvironment, and/or targeting metabolic checkpoints, are emerging. Most of the metabolic pathways described in this review also occur in normal cells, although they are frequently less active, making the determination of the right therapeutic window difficult. Thus, if we are able to distinguish particular requirements of cancer cells to take up and utilize or eliminate certain metabolites, specifically targeting these exchanges may provide more effective treatment strategies. Finally, as already described in several solid tumors, an in vitro examination of metabolic flux networks does not reflect what occurs in situ, in vivo, and in patients due mainly to the enormous plasticity and heterogeneity of their metabolism [219, 220, 202]. AML, in common with many tumors, is highly genetically heterogeneous and its metabolism should be directly studied in patients in situ.


Nitrogenous Waste in Birds and Reptiles: Uric Acid

Birds and reptiles have evolved the ability to convert toxic ammonia into uric acid or guanine rather than urea.

Learning Objectives

Compare the major byproduct of ammonia metabolism in mammals to that of birds and reptiles

Key Takeaways

Key Points

  • Nitrogenous wastes in the body tend to form toxic ammonia, which must be excreted.
  • Mammals such as humans excrete urea, while birds, reptiles, and some terrestrial invertebrates produce uric acid as waste.
  • Uricothelic organisms tend to excrete uric acid waste in the form of a white paste or powder.
  • Conversion of ammonia into uric acid is more energy intensive than the conversion of ammonia into urea.
  • Producing uric acid instead of urea is advantageous because it is less toxic and reduces water loss and the subsequent need for water.

Key Terms

  • urea: a water-soluble organic compound, CO(NH2)2, formed by the metabolism of proteins and excreted in the urine
  • guano: the excrement of seabirds, cave-dwelling bats, pinnipeds, or birds more generally
  • purine: any of a class of organic heterocyclic base containing fused pyrimidine and imidazole rings they are components of nucleic acids
  • xanthine: a precursor of uric acid found in many organs of the body
  • hypoxanthine: an intermediate in the biosynthesis of uric acid
  • uric acid: a bicyclic heterocyclic phenolic compound, formed in the body by the metabolism of protein and excreted in the urine

Nitrogenous Waste in Birds and Reptiles: Uric Acid

Of the four major macromolecules in biological systems, both proteins and nucleic acids contain nitrogen. During the catabolism, or breakdown, of nitrogen-containing macromolecules, carbon, hydrogen, and oxygen are extracted and stored in the form of carbohydrates and fats. Excess nitrogen is excreted from the body. Nitrogenous wastes tend to form toxic ammonia, which raises the pH of body fluids. The formation of ammonia itself requires energy in the form of ATP and large quantities of water to dilute it out of a biological system.

While aquatic animals can easily excrete ammonia into their watery surroundings, terrestrial animals have evolved special mechanisms to eliminate the toxic ammonia from their systems. The animals must detoxify ammonia by converting it into a relatively-nontoxic form such as urea or uric acid.

Nitrogen excretion: Nitrogenous waste is excreted in different forms by different species. These include (a) ammonia, (b) urea, and (c) uric acid.

Birds, reptiles, and most terrestrial arthropods, such as insects, are called uricothelic organisms because they convert toxic ammonia to uric acid or the closely-related compound guanine (guano), rather than urea. In contrast, mammals (including humans) produce urea from ammonia however, they also form some uric acid during the breakdown of nucleic acids. In this case, uric acid is excreted in urine instead of in feces, as is done in birds and reptiles.

Uric acid is a compound similar to purines found in nucleic acids. It is water insoluble and tends to form a white paste or powder. The production of uric acid involves a complex metabolic pathway that is energetically costly in comparison to processing of other nitrogenous wastes such as urea (from the urea cycle) or ammonia however, it has the advantages of reducing water loss and, hence, reducing the need for water.

Uric acid is also less toxic than ammonia or urea. It contains four nitrogen atoms only a small amount of water is needed for its excretion. Out of solute, it precipitates and forms crystals. The enzyme xanthine oxidase makes uric acid from xanthine and hypoxanthine, which in turn are produced from other purines. Xanthine oxidase is a large enzyme whose active site consists of the metal, molybdenum, bound to sulfur and oxygen. Uric acid is released in hypoxic conditions.


The role of NO

NO is a highly reactive gas with high diffusion rates across membranes. Several molecules can derive from NO and are collectively called reactive nitrogen species (RNS) that comprise the radical NO·, its nitrosonium (NO + ), and ni-troxyl ions (NO – ). When NO is produced in conjunction with ROS, such as during plant–pathogen interactions, NO can react with the superoxide anion O2· – to generate peroxynitrite (ONOO – ). In animals, the generation of NO under infectious conditions is mainly due to an inducible nitric oxide synthase (iNOS), which catalyses the NADPH-dependent oxidation of l -arginine to l -citrulline and NO ( Stuehr et al., 2004). In plants several data suggest the existence of such an enzymatic activity, but we still do not know the enzyme involved in this process ( Besson-Bard et al., 2008 Bellin et al., 2013). Production of NO in plants has been suggested to depend on several routes that can be divided into oxidative and reductive routes. Oxidative routes include the enzymatic activities polyamine and hydroxylamine oxidation ( Tun et al., 2006 Ruemer et al., 2009). Accordingly, a copper amine oxidase was proposed to be involved in NO production in planta in response to ABA ( Wimalasekera et al., 2011). Reductive routes consist of nitrite reduction via mitochondrial electron transfer systems ( Modolo et al., 2005 Planchet et al., 2005 Gupta and Igamberdiev, 2011), a root-specific nitrite:NO-reductase (Ni-NOR), the peroxisomal xanthine oxidoreductase enzyme ( Stohr et al., 2001), and nitrate reductase (NR) ( Rockel et al., 2002 Moche et al., 2010). NO is involved in several physiological processes in plants, including germination, development, stomatal closure, and immunity where it was shown to be involved in the hypersensitive response (HR) and during compatible interactions ( Delledonne et al., 1998, 2001 Durner et al., 1998). The role of NO in plant–pathogen interactions has been reviewed recently ( Bellin et al., 2013). Here, we will only focus on aspects that link N nutrition and metabolism to NO production in plant–pathogen interactions.

In the context of plant–pathogen interactions, it seems that NR is an important source of NO. Modolo et al. (2005) were the first to show that NR activity is the major source of NO during the pathogenic interaction ArabidopsisP. syringae. Other reports showed that NR participates in NO accumulation in plant–pathogen interactions ( Asai and Yoshioka, 2009 Perchepied et al., 2010) or in response to elicitors ( Yamamoto-Katou et al., 2006). However, decreased HR in Arabidopsis plants treated with P. syringae pv. maculicola in NR-deficient plants was correlated to a lack of l -arginine and NO2, two important endogenous substrates for NO synthesis ( Modolo et al., 2006). Conversely, it was later shown that the increased susceptibility to P. syringae of the NR-deficient plants was independent of amino acid accumulation and was more likely to be due to a reduced ability of these mutants to synthesize NO ( Oliveira et al., 2009). Interestingly, the activity of NIA2-encoded Arabidopsis NR enzyme was shown to be up-regulated through phosphorylation by the MAP kinase MPK6 ( Wang et al., 2010), involved in biotic stress responses ( Pitzschke et al., 2009) however, the role of this regulation during plant–pathogen interactions remains to be investigated.

Interestingly the nutrition of the plant can have an effect on NO production. Tobacco grown with nitrate was found to produce more NO than tobacco grown on ammonium when plants were inoculated with the pathogenic bacterium P. syringae pv. tabaci or the incompatible bacterium P. syringae pv. phaseolicola ( Gupta et al., 2013). The authors showed that NO accumulation was associated with increased resistance to the pathogens. Conversely, in soybean cotyledons, no difference was observed in NO production whether the plants were grown with nitrate or ammonium ( Galatro et al., 2013). Pathogens can contribute to the scavenging of NO. For instance, the flavohaemoglobin HmpX from the pathogenic bacterium D. dadantii was shown to contribute to the reduction of NO during HR ( Boccara et al., 2005). Thus NO is a pivotal element in plant–pathogen interactions, and its production and turnover are strongly linked to N metabolism.


Biological Nitrogen Fixation

Nitrogen is arguably the most important nutrient required by plants. However, the availability of nitrogen is limited in many soils and although the earth's atmosphere consists of 78.1% nitrogen gas (N2) plants are unable to use this form of nitrogen. To compensate , modern agriculture has been highly reliant on industrial nitrogen fertilizers to achieve maximum crop productivity. However, a great deal of fossil fuel is required for the production and delivery of nitrogen fertilizer. Moreover carbon dioxide (CO2) which is released during fossil fuel combustion contributes to the greenhouse effect and run off of nitrate leads to eutrophication of the waterways. Biological nitrogen fixation is an alternative to nitrogen fertilizer. It is carried out by prokaryotes using an enzyme complex called nitrogenase and results in atmospheric N2 being reduced into a form of nitrogen diazotrophic organisms and plants are able to use (ammonia). It is this process and its major players which will be discussed in this book.

Biological Nitrogen Fixation is a comprehensive two volume work bringing together both review and original research articles on key topics in nitrogen fixation. Chapters across both volumes emphasize molecular techniques and advanced biochemical analysis approaches applicable to various aspects of biological nitrogen fixation.

Volume 1 explores the chemistry and biochemistry of nitrogenases, nif gene regulation, the taxonomy, evolution, and genomics of nitrogen fixing organisms, as well as their physiology and metabolism.

Volume 2 covers the symbiotic interaction of nitrogen fixing organisms with their host plants, including nodulation and symbiotic nitrogen fixation, plant and microbial "omics", cyanobacteria, diazotrophs and non-legumes, field studies and inoculum preparation, as well as nitrogen fixation and cereals.

Covering the full breadth of current nitrogen fixation research and expanding it towards future advances in the field, Biological Nitrogen Fixation will be a one-stop reference for microbial ecologists and environmental microbiologists as well as plant and agricultural researchers working on crop sustainability.