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Chapter 13. Geomicrobially Important Interactions with Nitrogen

Chapter 13. Geomicrobially Important Interactions with Nitrogen

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TABLE 13.1

Abundance of Nitrogen in the Biosphere

Biosphere Compartment

Form of Nitrogen




Organic N

NH+4, NO −2, NO −3

Organic N

NH+4, NO −2, NO −3

Total N

Total N

Total N




Living biomass

Estimated Quantity

of Nitrogen (kg)

3.9 × 1018

9 × 1014


8 × 1014

1.4 × 1014

4 × 1017

1.9 × 1020

1.3 × 1013

Source: Extracted from Fenchel T, Blackburn TH, Bacteria and Mineral Cycling,

Academic Press, London, 1979; Brock TD, Madigan MT, Biology of

Microorganisms. 5th ed., Prentice-Hall, Englewood Cliffs, NJ, 1988.

remains). The first step in the recycling process is ammonification, in which the organic nitrogen is

transformed into ammonia. An example of ammonification is the deamination of amino acids:










The NH3 reacts with water to form ammonium hydroxide, which dissociates as follows:

NH 3 ϩ Η 2Ο → NH 4ΟΗ → NHϩ4 ϩ ΟΗϪ


In the laboratory it is commonly observed that when heterotrophic bacteria grow in proteinaceous

medium, such as nutrient broth consisting of peptone and beef extract, in which the organic nitrogen

serves as the source of energy, carbon, and nitrogen, the pH rises over time owing to the liberation

of ammonia and its hydrolysis to ammonium ion. Indeed, ammonification is always an essential first

step when an amino compound such as an amino acid serves as an energy source.

Ammonia is also formed as a result of urea hydrolysis catalyzed by the enzyme urease.

NH 2CONH 2 ϩ 2H 2Ο → 2NHϩ4 ϩ CΟ 32Ϫ


Urea is a nitrogen waste product excreted in the urine of many mammals. Although urease is produced by a variety of prokaryotic and eukaryotic microbes, a few prokaryotic soil microbes, for

example, Bacillus pasteurii and B. freudenreichii, seem to be specialists in degrading urea. They

prefer to grow at an alkaline pH such as that generated when urea is hydrolyzed (see Alexander,


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Geomicrobially Important Interactions with Nitrogen



Plants can readily assimilate ammonia. But the ammonia produced in ammonification in aqueous

systems at neutral pH exists as a positively charged ammonium ion (NH+4) owing to protonation. It is

adsorbed by clays and then not readily available to plants. Thus it is important that ammonia be converted into an anionic nitrogen species, which is not readily adsorbed to the negatively charged clays

and is thus more readily available to plants. Nitrifying bacteria play a central role in this conversion.

The majority of nitrifying bacteria are autotrophs and can be divided into two groups; one includes

those bacteria that oxidize ammonia to nitrous acid (e.g., Nitrosomonas, Nitrosocystis)

s and the

second includes those bacteria that oxidize nitrite to nitrate (e.g., Nitrobacter,

r Nitrococcus).

s All the

members of these two groups of nitrifying bacteria are obligate autotrophs except for Nitrobacter

winogradskyi strains, which appear to be facultative. They are all strict aerobes. Representatives

are found in soil, freshwater, and seawater (for further characterization see Paul and Clark, 1996;

Balows, 1992).

Although ammonification is the major source of ammonia in soil and sediments, a special anaerobic respiratory process may also be a significant source of ammonia in some environments. In this

process, nitrate is reduced to ammonia via nitrite (Jørgensen and Sørensen, 1985, 1988; Binnerup

et al., 1992). The overall process can be summarized in the equation

NOϪ3 ϩ 8(H) ϩ Ηϩ → NH 3 ϩ 3H 2Ο


This process is known as nitrate ammonification and can be carried on by a number of different facultative and strictly anaerobic bacteria (see Cruz-García et al., 2007; Dannenberg et al.,

1992; Sørensen, 1987; review by Ehrlich, 1993, pp. 232–233). A variety of organic compounds, H2,

and inorganic sulfur compounds can serve as electron donors in this reaction (Dannenberg et al.,




Oxidation of ammonia by ammonia-oxidizing bacteria involves hydroxylamine (NH2OH) as an

intermediate (see review by Wood, 1988). The formation of hydroxylamine is catalyzed by an


NH 3 ϩ 0.5Ο 2 → NH 2ΟΗ (∆G° ϭϩ0.77 kcal; ϩ3.21 kJ)

This reaction does not yield biochemically useful energy. Indeed, it is slightly endothermic and proceeds in the direction of hydroxylamine because it is coupled to the oxidation of hydroxylamine to

nitrous acid, which is strongly exothermic. The overall reaction of the oxidation of hydroxylamine

to HNO2 can be summarized as

NH 2OH ϩ O 2 → HΝO 2 ϩ H 2O (∆G° ϭ Ϫ62.42 kcal; Ϫ260.9 kJ)


It is Reaction 13.6 from which chemoautotrophic ammonia oxidizers obtain their energy, by using

chemiosmotic coupling, that is, oxidative phosphorylation. The conversion of hydroxylamine to

nitrous acid involves some intermediate steps (Hooper, 1984).

The enzyme that catalyzes ammonia oxidation (Reaction 13.5) is a nonspecific oxygenase. It can

also catalyze the oxygenation of methane to methanol (Jones and Morita, 1983).

CH 4 ϩ 0.5Ο 2 → CH 3ΟΗ (∆G° ϭ Ϫ29.74 kcal; Ϫ124.3 kJ)

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Under standard conditions, this reaction is thermodynamically more favorable than Reaction 13.5.

This does not mean, however, that ammonia oxidizers can grow on methane. They lack the ability

to oxidize methanol.

Ammonia oxidizers can also form some NO and N2O in side reactions of ammonia oxidation

under oxygen limitation in which nitrite replaces oxygen as terminal electron acceptor (see discussion by Knowles, 1985; Bock et al., 1991; Davidson, 1993). This is an important observation

because it means that biogenically formed N2O and NO are not solely the result of denitrification

(see Section 13.2.7).



The nitrite oxidizers convert nitrite to nitrate:

NOϪ2 ϩ 0.5Ο 2 → ΝΟϪ3 (∆G° ϭ Ϫ18.18 kcal; Ϫ76.0 kJ)

They obtain useful energy from this process by coupling it chemiosmotically to ATP generation

(Aleem and Sewell, 1984; Wood, 1988).



Ammonia is also converted to nitrate by certain heterotrophic microorganisms, but the process

is probably of minor importance in nature in most instances. Rates of heterotrophic nitrification

measured under laboratory conditions so far are significantly slower than those of autotrophic

nitrification. The organisms capable of heterotrophic nitrification include both bacteria, such as

Arthrobacterr sp., and fungi, such as Aspergillus flavus. They gain no energy from the conversion.

The pathway from ammonia to nitrate may involve intermediates such as hydroxylamine, nitrite,

and 1-nitrosoethanol in the case of bacteria; and 3-nitropropionic acid in the case of fungi (see

Alexander, 1977; Paul and Clark, 1996).



Within the last decade, some Planctomycetes (domain Bacteria) have been found to be involved in

the anaerobic oxidation of ammonium (NH+4) with nitrite (NO−2 ) to form dinitrogen (N2 ) (see van

de Graaf et al., 1995; and reviews by Jetten et al., 2003; Kuenen and Jetten, 2001; Strous and Jetten,

2004). An equation summarizing the overall reaction is as follows:

NHϩ4 ϩ ΝΟϪ2 → Ν 2 ϩ 2Η 2Ο (∆G° ϭ Ϫ85.51 kcal; Ϫ357.79 kJ)


Hydrazine (N2H4) and hydroxylamine (NH2OH) are intermediates. Acetylene and methanol have

been found to inhibit different reactions in anammox (Jensen et al., 2007). Anammox was first

discovered in an anaerobic sewage treatment process but has since been demonstrated to occur in

nature in both freshwater and marine environments (Jetten et al., 2003; Trimmer et al., 2003; Tal

et al., 2005). In the oceans it may account for 50% of the loss of fixed nitrogen (Dalsgaard et al.,

2005). The free energy from this reaction (Equation 13.9) supports CO2 fixation. The anammox

reaction appears to be carried out in a special intracytoplasmic organelle in active bacteria. The

organelle is called an anammoxosome. It is bounded by a membrane that contains ladderane lipids

(see van Niftrik et al., 2004). The bacteria capable of the anammox reaction belong to the phylum Planctomycetes. Candidatus Brocadia anammoxidans is one member that has been described

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Geomicrobially Important Interactions with Nitrogen


(Strous et al., 1999; Kuenen and Jetten, 2001). Two others are Candidatus Kuenenia stuttgartiensis

and Candidatus Scalindua sorokinii (Jetten et al., 2003).


Nitrate, nitrite, and nitrous and nitric oxides can serve as electron acceptors in microbial respiration,

usually under anaerobic conditions. The transformation of nitrate to nitrite as an anaerobic respiratory process is called dissimilatory nitrate reduction, and the reduction of nitrate to nitric oxide

(NO), nitrous oxide (N2O), and dinitrogen is called denitrification. Assimilatory nitrate reduction

is the first step in a process in which nitrate is reduced to ammonia for the purpose of assimilation.

Only as much nitrate is consumed in this process as is needed for assimilation. It is not a form of respiration and is performed by many organisms that cannot use nitrate for respiration. Some nitraterespiring bacteria are capable of only nitrate reduction, lacking the enzymes for reduction of nitrite

to dinitrogen, whereas others are capable of reducing nitrite to ammonia instead of dinitrogen in a

process that has been called nitrate ammonification by Sørensen (1987) (see also Section 13.2.6).

All nitrate respiratory processes have been found to operate to varying degrees in terrestrial, freshwater, and marine environments and represent an important part of the nitrogen cycle favored by

anaerobic conditions.

Nitrate reduction is described by the half-reaction

NOϪ3 ϩ 2Hϩ ϩ 2e → NOϪ2 ϩ H 2O


The electron donor may be any one of a variety of organic metabolites or reduced forms of sulfur

such as H2S or S0. The enzyme catalyzing reaction 13.10 is called nitrate reductase and is an iron

molybdoprotein. It is not only capable of catalyzing nitrate reduction but may also catalyze reduction of ferric to ferrous iron (see Chapter 16) and reduction of chlorate to chlorite. Nitrate can competitively affect ferric iron reduction by nitrate reductase (Ottow, 1969).

The reduction of nitrite to dinitrogen is illustrated by the following series of half-reactions, with

organic metabolites or reduced sulfur acting as electron donor:

NOϪ2 ϩ 2Hϩ ϩ e → NO ϩ H 2O


2NO ϩ 2Hϩ ϩ 2e → N 2O ϩ H 2O


N 2O ϩ 2Hϩ ϩ 2e → N 2 ϩ H 2O


The reduction of nitrite to ammonia may be summarized by the equation

NOϪ2 ϩ 7Hϩ ϩ 6e → NH 3 ϩ 2H 2O


The electron donor may be one of a variety of organic metabolites.

Although it was previously believed that these reactions can only occur at low oxygen tension or in the absence of oxygen, evidence now indicates that in some cases organisms such as

Thiosphaera pantotropha can perform the reactions at near-normal oxygen tension (Robertson and

Kuenen, 1984a,b; but see also disagreement by Thomsen et al., 1993). This organism can actually

use oxygen and nitrate simultaneously as terminal electron acceptors. The explanation is that in Tsa.

pantotropha the enzymes of denitrification are produced aerobically as well as anaerobically, whereas in

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oxygen-sensitive denitrifiers they are produced only at low oxygen tension or anaerobically. Nitrate

reductase in Tsa. pantotropha is constitutive, whereas in many anaerobic denitrifiers it is inducible. Moreover, nitrate reductase in Tsa. pantotropha is not inactivated by oxygen, unlike in some

anaerobic denitrifiers. Finally, oxygen does not repress the formation of the denitrifying enzymes

in Tsa. pantotropha, whereas it does in some anaerobic denitrifiers. Aerobic denitrification in Tsa.

pantotropha appears to be linked to heterotrophic nitrification (Robertson et al., 1988; Robertson

and Kuenen, 1990). The organism seems to use denitrification as a means of disposing of excess

reducing power because its cytochrome system is insufficient for this purpose. Oxygen tolerance in

denitrification has also been observed with some other bacteria (Hochstein et al., 1984; Davies et

al., 1989; Bonin et al., 1989).

For a more complete discussion of denitrification, the reader is referred to a monograph by Payne

(1981) and a review article by the same author (Payne, 1983).


If nature had not provided for microbial nitrogen fi xation to reverse the effect of microbial depletion

of fixed nitrogen from soil or water as volatile nitrogen oxides or dinitrogen as a result of denitrification and anammox, life on Earth would not have long continued after the processes of denitrification

and anammox first evolved. Nitrogen fixation is dependent on a special enzyme, nitrogenase, which

is found only in prokaryotic organisms, including aerobic and anaerobic photosynthetic and nonphotosynthetic Bacteria and Archaea. Nitrogenase is an oxygen-sensitive enzyme, usually a combination of an iron protein and a molybdoprotein (Eady and Postgate, 1974; Orme-Johnson, 1992), but

in some cases (e.g., Azotobacter chroococcum)

m may also be a combination of an iron protein and a

vanadoprotein (Robson et al., 1986; Eady et al., 1987), and in another case ((A. vinelandii) may be a

combination of two iron proteins (Chiswell et al., 1988). Nitrogenase catalyzes the reaction

N 2 ϩ 6Ηϩ ϩ 6e → 2NH 3


The enzyme is not specific for dinitrogen. It can also catalyze the reduction of acetylene (CHCH), as

well as of hydrogen cyanide (HCN), cyanogen (NCCN), hydrogen azide (HN3), hydrogen thiocyanate (HCNS), protons (H+), carbon monoxide (CO), and some other compounds (Smith, 1983).

The reducing power (the term “6e” in Equation 13.15) needed for dinitrogen reduction is provided by reduced ferredoxin. Reduced ferredoxin can be formed in a reaction in which pyruvate is

oxidatively decarboxylated (Lehninger, 1975),

CH 3COCOOH ϩ ΝΑDϩ ϩ CoASH → CH3CO ∼ SCoA ϩ CO2 ϩ ΝΑDH ϩ Hϩ


NADH ϩ (ferredoxin)ox → NADϩ ϩ (ferredoxin)red ϩ Ηϩ


In phototrophs, the reduced ferrodoxin is produced as part of the photophosphorylation mechanism

(see Chapter 6).

Nitrogen fixation is a very energy-intensive reaction, consuming as many as 16 moles of ATP in

the reduction of 1 mole of dinitrogen to ammonia (Newton and Burgess, 1983).

Nitrogen fixation may proceed symbiotically or nonsymbiotically. Symbiotic nitrogen fi xation

requires that the nitrogen-fixing bacterium associates with a specific host plant (e.g., a legume), one

of several nonleguminous angiosperms, the water fern Azolla, fungi (certain lichens), or, in rare

cases, with an animal host to carry out nitrogen fixation. Even then, dinitrogen will be fixed only

if the fixed-nitrogen level in the surrounding environment of the host plant is low or the diet of the

animal host is nitrogen-deficient. In some plants (legumes or alder), the nitrogen fixer may be localized in the cells of the cortical root tissue that are transformed into nodules. Invasion of the plant

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Geomicrobially Important Interactions with Nitrogen


tissue may have occurred via root hairs. In some other plants, the nitrogen fixers may be localized in

special leaf structures (e.g., in Azolla). In animals, the nitrogen fixer may be found to be a member

of the flora of the digestive tract (Knowles, 1978). The plant host in symbiotic nitrogen fixation provides the energy source required by the nitrogen fixer for generating ATP. The energy source may

take the form of compounds such as succinate, malate, and fumarate (Paul and Clark, 1996). The

plant host also provides an environment in which access to oxygen is controlled so that nitrogenase

in the nitrogen fixer is not inactivated. In root nodules of legumes, leghemoglobin is involved in the

control of oxygen. The nitrogen fixer shares the ammonia that it forms from dinitrogen with its plant

or animal host.

Symbiotic nitrogen-fixing bacteria include members of the genera Rhizobium, Bradyrhizobium,

Frankia, and Anabaena. Some strains of B. japonicum have been found to be able to grow autotrophically on hydrogen as energy source because they possess uptake hydrogenase. They can couple hydrogen oxidation to ATP synthesis, which they can use in CO2 assimilation via the ribulose

bisphosphate carboxylase/oxidase system. In nitrogen fixation, the ability to couple hydrogen oxidation to ATP synthesis may represent an energy recovery system because nitrogenase generates a

significant amount of hydrogen during nitrogen fixation, the energy content of which would otherwise be lost to the system.

About 30 years ago, a special symbiotic nitrogen-fixing relationship was discovered in Brazil

between certain cereal grasses, such as maize, and nitrogen-fixing spirilla such as Azospirillum

lipoferum (Von Bülow and Döbereiner, 1975; Day et al., 1975; Smith et al., 1976). In these symbioses, the nitrogen-fixing bacterium does not invade the host plant roots or any other part of the

plant but lives in close association with the roots in the rhizosphere. Apparently, the plants excrete

compounds via their roots that the nitrogen fixer can use as energy sources and that enable it to fix

nitrogen that it can share with the plant if the soil is otherwise deficient in fixed nitrogen.

In nonsymbiotic nitrogen fi xation, the active organisms are free-living in soil or water and fix

nitrogen if fixed nitrogen is limiting. Their nitrogenase is not distinctly different from that of symbiotic nitrogen fixers. Unlike the symbiotic nitrogen fixers, aerobic nonsymbiotic nitrogen fixers

appear to be able to maintain an intracellular environment in which nitrogenase is not inactivated

by oxidizing conditions. The capacity for nonsymbiotic nitrogen fixation is widespread among

prokaryotes. The best-known and the most efficient examples include the aerobes Azotobacter

and Beijerinckia and the anaerobe Clostridium pasteurianum, but many other aerobic and anaerobic genera include species with nitrogen-fixing capacity, even some photo- and chemolithotrophs.

Most of the nitrogen fixers are active only at environmental pH values between 5 and 9, but some

strains of the acidophile Acidithiobacillus ferrooxidans have been shown to fix nitrogen at a pH

as low as 2.5.

For a more detailed discussion of nitrogen fixation, the reader is referred to Alexander (1984),

Broughton (1983), Newton and Orme-Johnson (1980), and Quispel (1974).


Owing to their special capacities of transforming inorganic compounds, which plants and animals lack, microbes, especially prokaryotes and certain fungi, play a central role in the nitrogen

cycle (Figure 13.1). Many reactions of the cycle are entirely dependent on them; nitrogen fixation is

restricted to prokaryotes. The direction of transformations in the cycle is determined by environmental conditions, especially the availability of oxygen, but also by the supply of particular nitrogen compounds. Anaerobic conditions may encourage denitrification and anammox and thus cause

nitrogen limitation unless the process is counteracted by anaerobic nitrogen fixation. Limitations

in the supply of organic or ammonia nitrogen affect the availability of nitrate. Availability of fixed

nitrogen has been viewed as a growth-limiting factor in the marine environment but infrequently in

unpolluted freshwater, in which phosphate is more likely to limit productivity. Fixed nitrogen can

be a limiting factor in soil, especially in agriculturally exploited soils.

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Organic nitrogen

Guano, etc.
















FIGURE 13.1 The nitrogen cycle. (A) Ammonification, aerobic and anaerobic; (B) autotrophic nitrification,

strictly aerobic; (C) nitrate assimilation, aerobic and anaerobic; (D) nitrate reduction, usually anaerobic, but

see (D + E); (D + E) denitrification, usually anaerobic but sometimes aerobic, see text; (F) nitrogen fixation,

aerobic and anaerobic; (D + G) nitrate ammonification, anaerobic; and (H) anammox, anaerobic.



Nitrogen is essential to all forms of life. It is assimilated by cells in the form of ammonia or nitrate,

and in some instances as amino-nitrogen, mostly in the form of one or more amino acids. It is

released from organic combination in the form of ammonia. The latter process is called ammonification. It occurs both aerobically and anaerobically. Ammonia is an energy-rich compound and can

be oxidized to nitrate by way of nitrite by some aerobic, autotrophic bacteria (nitrifiers). It can also

be converted to nitrate by some heterotrophic bacteria and some fungi in a nonenergy yielding process, but this is much less common. The conversion of ammonia to nitrate is important in soil and

sediments because negatively charged clay particles can adsorb ammonia, making it unavailable

to plants. Under reducing conditions, nitrate can be transformed by anaerobic respiration to nitrite,

nitric and nitrous oxide, and dinitrogen, or ammonia by appropriate bacteria. In soil, the reduction

of nitrate to dinitrogen can have the effect of lowering its fertility, as can the anaerobic oxidation of

ammonia to dinitrogen by nitrite (anammox). Depletion of soil nitrogen through dinitrogen evolution can, however, be reversed by symbiotic and nonsymbiotic nitrogen-fixation by bacteria, which

are able to reduce dinitrogen to ammonia. These various biological interactions are part of a cycle

that is essential to the sustenance of life on Earth.


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14 Geomicrobial

with Arsenic and Antimony


Although arsenic and antimony are generally toxic to life, some microorganisms exist that can

metabolize some forms of these elements. Some can use arsenite or stibnite as partial or sole

energy sources whereas others can use arsenate as terminal electron acceptors. Still other microbes

can metabolize arsenic and antimony compounds to detoxify them. These reactions are important

from a geomicrobial standpoint because they indicate that some microbes contribute to arsenic

and antimony mobilization or immobilization in the environment and play a role in arsenic and

antimony cycles.





Arsenic is widely distributed in the upper crust of the Earth, mostly at very low concentrations. Its average abundance in igneous rocks has been estimated to be of the order of 5 g tn−1

(Carapella, 1972). It rarely occurs in elemental form. More often it is found to be combined with

sulfur, as in orpiment (As2S3), realgar (As2S2 or AsS), or arsenopyrite (FeAsS); with selenium,

as in As2Se3; with tellurium, as in As2Te; or as a sulfosalt, as in enargite (Cu3AsS4). It is also

found in arsenides of heavy metals such as iron (loellingite, FeAs2), copper (domeykite, Cu3As),

nickel (nicolite, NiAs), and cobalt (Co2As). Sometimes it occurs in the form of arsenite minerals

(arsenolite or claudetite, As2O3) or arsenate minerals (erythrite, Co3(AsO4)2∙8H 2O; scorodite,

FeAsO4 ∙ 2H 2O; olivenite, Cu2(AsO4)(OH)). Arsenopyrite is the most common and widespread

mineral form of arsenic, but orpiment and realgar are also fairly common. The ultimate source

of arsenic on the Earth’s surface is igneous activity. On weathering of arsenic-containing rocks,

which may harbor as much as 1.8 ppm of the element, the arsenic is dispersed through the upper

lithosphere and the hydrosphere.

Arsenic concentration in soil may range from 0.1 to more than 1000 ppm. The average concentration in seawater has been reported to be 3.7 µg L−1 and in freshwater, 1.5–53 ng m−3 (Bowen,

1979). However, in groundwater in the Munshiganj District of Bangladesh, the As concentration

approaches a maximum of 640 mg m−3 at a depth of 30–40 m but decreases to 58 mg m−3 at 107 m

(Swartz et al., 2004). Some living organisms may concentrate arsenic many fold over its level in the

environment. For example, some algae have been found to accumulate arsenic 200–3000 times in

excess of its concentration in the growth medium (Lunde, 1973). Humans may artificially raise the

arsenic concentrations in soil and water through the introduction of sodium arsenite (NaAsO3) or

cacodylic acid ((CH3)2AsO ∙ OH) as herbicides.


In nature, arsenic is usually encountered in the oxidation states of 0, +3, and +5. Its coordination

numbers are in the range of 3–6 (Cullen and Reimer, 1989). Except for the oxidation states of As

in arsenate and arsenite, the oxidation state of other compounds, whether organic or inorganic, is


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often unclear and depends on its definition (Cullen and Reimer, 1989, p. 715). According to Cullen

and Reimer (1989, p. 715), arsenious acid and its salts in aqueous solution exist in the ortho form

(H3AsO3) but not in the meta (HAsO2) form. Environmentally, arsenite is more mobile than arsenate,

but it can be significantly adsorbed under certain conditions. This is because of the tendency of

arsenate to become strongly adsorbed to mineral surfaces such as those of ferrihydrite. However,

an ability of ferrihydrite and schwertmannite to adsorb both As(V) and As(III) has been observed

by Raven et al. (1998) and Carlson et al. (2002). The ability of disordered mackinawite (FeS),

which can be formed in sediments where bacterial sulfate reduction occurs, to adsorb As(V) and

As(III) has been reported by Wolthers et al. (2005). As(III) adsorption by amorphous iron oxide

and goethite has also been studied by Dixit and Hering (2003, 2006), who noted preferential uptake

of As(V) in a pH range of 5–6 and As(III) in a pH range of 7–8. They also noted that with goethite,

current single-sorbate models were able to predict the adsorption of Fe(II) and As(III) more satisfactorily than double-sorbate models. Overall, because of the lesser mobility of arsenate compared

with arsenite under common environmental conditions, reduction of arsenate to arsenite in the

environment, whether chemical or biological, leads to an increase in arsenic toxicity (Cullen and

Reimer, 1989).



Arsenic compounds are toxic for most living organisms. Arsenite (AsO33−) has been shown to inhibit

dehydrogenases such as pyruvate, α-ketoglutarate-, and dihydrolipoate-dehydrogenases (Mahler

and Cordes, 1966). Arsenate (AsO43−) uncouples oxidative phosphorylation, that is, it inhibits chemiosmotic ATP synthesis (Da Costa, 1972).

Both the uptake and the inhibitory effect of arsenate on metabolism can be modified by

phosphate (Button et al., 1973; Da Costa, 1971, 1972). This is because of the existence of a common transport mechanism for phosphate and arsenate in the membranes of some organisms.

However, a separate transport mechanism for phosphate may also exist (Bennett and Malamy,

1970). In the latter case, phosphate uptake does not affect arsenate uptake, nor does arsenate

uptake affect phosphate uptake. In one reported case of a fungus, Cladosporium herbarium,

arsenite toxicity could also be ameliorated by the presence of phosphate. In that instance,

prior oxidation of the arsenite to arsenate by the fungus appeared to be the basis for the effect

(Da Costa, 1972). In growing cultures of Candida humicola, phosphate can inhibit the formation

of trimethylarsine from arsenate, arsenite, and monomethylarsonate, but not from dimethylarsinate (Cox and Alexander, 1973). In similar cultures, phosphite can suppress the formation of trimethylarsine from monomethylarsonate, but not from arsenate or dimethylarsinate.

Hypophosphite can cause temporary inhibition of the conversion of arsenate, monomethylarsonate, and dimethylarsinate (Cox and Alexander, 1973). High antimonite concentrations

lower the rate of conversion of arsenate to trimethylarsine by resting cells of C. humicola (Cox and

Alexander, 1973).

Bacteria can develop genetically determined resistance to arsenic (Ji and Silver, 1992; Ji et al.,

1993). The gene locus for this resistance may reside on a plasmid as, for example, in Staphylococcus

aureus (Dyke et al., 1970) and Escherichia coli (Hedges and Baumberg, 1973). The mechanism of

resistance in these bacterial species is a special pumping mechanism that expels the arsenic taken

up as arsenate by the cells (Silver and Keach, 1982). It involves intracellular reduction of arsenate to arsenite followed by efflux of the arsenite promoted by an oxyanion-translocating ATPase

(Broeer et al., 1993; Ji et al., 1993). Some of the resistant organisms have the capacity to oxidize

reduced forms of arsenic to arsenate and others to reduce oxidized forms (see Sections 14.2.4 and

14.2.6). In Shewanella sp. strain ANA-3, the traits of arsenate resistance and arsenate respiration

are encoded in two distinct genetic loci, operons ars and arrs, respectively (Saltikov and Newman,

2003; Saltikov et al., 2003; see also Section 14.2.7).

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