Is there a fundamental reason that plants cannot fix their own nitrogen?

Is there a fundamental reason that plants cannot fix their own nitrogen?

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Plants must have nitrogen to grow. According to the answer to this question, there are no plants that can fix their own nitrogen (without the help of bacteria).

Plants get their nitrogen in the form of nitrates (NO3-) or ammonia (NH4+). Nitrates and ammonia get into the soil through

  • lightning (causing N2 and O2 to combine to form NO which then reacts with atmospheric water and is brought to earth by rain)
  • nitrogen-fixing bacteria associated with leguminous plants
  • fixed nitrogen from dead plants/animals (ammonification), urine, etc.
  • ammonifying and other bacteria

The answer also states that it seems to be possible to engineer plants to be able to fix nitrogen.

So one could think plants could have evolved to be able to fix nitrogen, but did not. Maybe it just did not happen based on it's probability to happen.
Is there any other explanation?

Maybe the ability to fix nitrogen would have a harmful effect on plants? Or fixation requires a complex set of related mutations? Are there enough areas with soil depleted of nitrates to exert a selection pressure for nitrogen-fixing plants?

That is an interesting question. But rises some line of thought by analogy, e.g. why people won't evolve to produce their own ATP without mitochondria? (I mean of course Eukaryotas in general). I don't see any reason why multicellular organism cannot incorporate bacterial functionality either as happened with mitochondria, or as just separate cell type, that will provide desirable enzymatic reactions.

The answer, no matter how unsatisfactory, might be simple: because it so happened. We definitely can create experimental conditions lacking N-fixing bacteria as well as varying amounts of nitrates, nitrites etc. And then produce and screen line of mutants for ability to fix nitrogen without symbiotic partners.

The most obvious answer to your question is that the enzyme responsible for nitrogen fixation is inactivated by oxygen. So photosynthesis, which produces oxygen, rules out nitrogen fixation, and, as you know, plants photosynthesise.

However, they could have evolved specialized cells for nitrogen fixation, which do not photosynthesise. In fact, some colonial cyanobacteria, which photosynthesise, form such specialized cells called heterocysts.

So why haven't plants evolved such specialized cells? I think a fundamental answer to this question would invoke the evolutionary theory of specialization which says that specialization in one thing at the cost of the ability to do something else is selected for when the trade-off is convex, i.e. when you can be really good at only one thing, and being able to do both implies you are actually quite bad at both.

I hope this sets you off to find the fundamental reason you are looking for.

I believe that evolution is driven by the needs of the organism to adapt not by the need to adapt for human beings consumption. meaning that some nitrogen gets back to the soil through the effect of nature e.g. fixation of nitrogen during storms and when fecal matter decomposes in soil and these I assume provide enough nitrogen for normal plant growth and so the plant is not "forced" to adapt to say a deficiency in nitrogen but for humans I assume because of agriculture the land gets "exhausted" and so we need fertilisers but plants have not evolved becuase I assume there was no pressing need for it. That is my own thinking assuming I understood your question

Soil Inoculants

Soil biology is important for keeping agricultural systems healthy and productive. Living soil is complex. It includes creatures that cannot be seen with the naked eye, such as bacteria, fungi, actinomycetes, protozoa and nematodes, as well as creatures such as insects and earthworms. This community of organisms is bound together in a food web that affects the soil's chemical and physical properties. We care about these properties because they also affect plant growth and health.

Practices such as adding manures or composts to soil, planting cover crops and rotating crops are all aimed at rebuilding and maintaining soil organic matter, recycling and retaining nutrients, and decreasing soil diseases. These practices are usually associated with increased microbial biomass and increased soil organism diversity.

A healthy soil can contain billions of bacteria, fungi and other microorganisms in one teaspoon. Depending on soil conditions, the populations of these different microorganisms rise and fall. Some microbial populations increase quickly when fresh cover crops or other plant residues are added to the soil. For example, some microbes are able to use the readily available sources of carbon from fresh plant residues like humans use carbohydrates. These microbes decrease as the carbon sources are used up, causing other microbes that break down the less available sources of carbon like cellulose and lignin to increase. The point is that there are many native microorganisms in the soil that respond quickly when conditions are favorable for their growth.


In their native form, plants constitute a remarkable feat of metabolic engineering. Not only does their energy derive entirely from the sun and their carbon from CO2, but they can defend themselves from pests and predators without the benefit of mobility they participate in complex symbioses, in part by tailoring the composition of their epi- and endophytic microbial communities, and they can survive extremes of temperature and nutrient and water availability. What more could we ask of plants?

A great deal, it turns out. Both conventional breeding and modern metabolic engineering have been used to boost productivity and to enhance fitness (for example, by increased resistance to pests, herbicides, and climatic extremes) [1]. In addition, new areas of application have been introduced that would have seemed like science fiction only a few decades ago, including the use of plants to produce vaccines, bioplastics, and derivatives of complex natural product drugs [2]–[4]. Many of these more recent engineering goals could not have been accomplished by fine-tuning endogenous host metabolism instead, they required the installation of new metabolic pathways from other plants or bacteria. Adding new nodes to a plant metabolic network is a difficult task that will benefit from advances in targeted genome modification, tissue-, cell-, and organelle-specific gene expression, and the controlled expression of multi-gene pathways [5]–[7].

In this essay, we highlight recent progress in, and the near-term potential of, four long-standing grand challenges in plant metabolic engineering: two deal with important applications in food and energy, while the remaining two are of general utility in improving plant fitness, and in principle would be useful for improving plants as a chassis for other metabolic engineering efforts. Nature never intended plants to be grown as crops on an industrial scale, nor did plants evolve solely for human nourishment. Plants are not naturally inclined to give up their structural oligosaccharides in ready-to-eat form in the service of providing green energy. Although each of these challenges has been recognized for decades and important advances have been made [8]–[10], solutions to them still lie far beyond our current capabilities. Nevertheless, the technologies developed to meet them will have myriad uses long before the problems themselves are solved. Techniques for using synthetic biology to make multiple deletions, additions, and other edits to plant genomes stand out as a particularly important set of enabling technologies for the challenges described below [11]. Finally, while we focus primarily on the technical aspects involved in developing these engineering efforts, we recognize that addressing societal acceptance, economic considerations, environmental impact, and long-term sustainability are also of critical importance for their successful implementation.


Biological nitrogen fixation was discovered by Jean-Baptiste Boussingault in 1838. [9] Later, in 1880, the process by which it happens was discovered by German agronomist Hermann Hellriegel and Hermann Wilfarth [de] [10] and was fully described by Dutch microbiologist Martinus Beijerinck. [11]

"The protracted investigations of the relation of plants to the acquisition of nitrogen begun by Saussure, Ville, Lawes and Gilbert and others culminated in the discover of symbiotic fixation by Hellriegel and Wilfarth in 1887." [12]

"Experiments by Bossingault in 1855 and Pugh, Gilbert & Lawes in 1887 had shown that nitrogen did not enter the plant directly. The discovery of the role of nitrogen fixing bacteria by Herman Hellriegel and Herman Wilfarth in 1886-8 would open a new era of soil science." [13]

In 1901 Beijerinck showed that azotobacter chroococcum was able to fix atmospheric nitrogen. This was the first species of the azotobacter genus, so-named by him. It is also the first known diazotroph, the species that use diatomic nitrogen as a step in the complete nitrogen cycle.

Biological nitrogen fixation (BNF) occurs when atmospheric nitrogen is converted to ammonia by a nitrogenase enzyme. [1] The overall reaction for BNF is:

The process is coupled to the hydrolysis of 16 equivalents of ATP and is accompanied by the co-formation of one equivalent of H
2 . [14] The conversion of N
2 into ammonia occurs at a metal cluster called FeMoco, an abbreviation for the iron-molybdenum cofactor. The mechanism proceeds via a series of protonation and reduction steps wherein the FeMoco active site hydrogenates the N
2 substrate. [15] In free-living diazotrophs, nitrogenase-generated ammonia is assimilated into glutamate through the glutamine synthetase/glutamate synthase pathway. The microbial nif genes required for nitrogen fixation are widely distributed in diverse environments. [16]

For example decomposing wood which has generally low content of nitrogen was shown to host diazotrophic community. [17] [18] Bacteria through fixation enrich wood substrate with nitrogen thus enabling deadwood decomposition by fungi. [19]

Nitrogenases are rapidly degraded by oxygen. For this reason, many bacteria cease production of the enzyme in the presence of oxygen. Many nitrogen-fixing organisms exist only in anaerobic conditions, respiring to draw down oxygen levels, or binding the oxygen with a protein such as leghemoglobin. [1]

Importance of nitrogen Edit

Atmospheric nitrogen is inaccessible to most organisms, [20] because its triple covalent bond is very strong. The nitrogen requirements for life is highly variable. [ clarification needed ] Considering atom acquisition, for every 100 atoms of carbon, roughly 2 to 20 atoms of nitrogen are assimilated. The atomic ratio of Carbon(C): Nitrogen(N): Phosphorus(P) observed on average in planktonic biomass was originally described by Alfred Redfield. [21] The Redfield Ratio, the stoichiometric relationship between C:N:P atoms, is 106:16:1. [21]

Nitrogenase Edit

The protein complex nitrogenase is responsible for catalyzing the reduction of nitrogen gas (N2) to ammonia (NH3). [22] In Cyanobacteria, this enzyme system is housed in a specialize cell called the heterocyst. [23] The production of the nitrogenase complex is genetically regulated, and the activity of the protein complex is dependent on ambient oxygen concentrations, and intra- and extracellular concentrations of ammonia and oxidized nitrogen species (nitrate and nitrite). [24] [25] [26] Additionally, the combined concentrations of both ammonium and nitrate are thought to inhibit NFix, specifically when intracellular concentrations of 2-oxoglutarate (2-OG) exceed a critical threshold. [27] The specialized heterocyst cell is necessary for the performance of nitrogenase as a result of its sensitivity to ambient oxygen. [28]

Nitrogenase consist of two proteins, a catalytic iron-dependent protein, commonly referred to as MoFe protein and a reducing iron-only protein (Fe protein). There are three different iron dependent proteins, molybdenum-dependent, vanadium-dependent, and iron-only with all three nitrogenase proteins variations containing an iron protein component. Molybdenum-dependent nitrogenase is the most commonly present nitrogenase. [22] The different types of nitrogenase can be determined by the specific iron protein component. [29] Nitrogenase is highly conserved, gene expression through DNA sequencing can distinguish which protein complex is present in the microorganism and potentially being express. Most frequently, the nifH gene is used to identify the presence of molybdenum-dependent nitrogenase followed by closely related nitrogenase reductases (component II) vnfH and anfH representing vanadium-dependent and iron-only nitrogenase, respectively. [30] In studying the ecology and evolution of nitrogen-fixing bacteria, the nifH gene is the biomarker most widely used. [31] nifH has two similar genes anfH and vnfH that also encode for the nitrogenase reductase component of the nitrogenase complex [32]

Microorganisms Edit

Diazotrophs are widespread within domain Bacteria including cyanobacteria (e.g. the highly significant Trichodesmium and Cyanothece), as well as green sulfur bacteria, Azotobacteraceae, rhizobia and Frankia. Several obligately anaerobic bacteria fix nitrogen including many (but not all) Clostridium spp. Some archaea also fix nitrogen, including several methanogenic taxa, which are significant contributors to nitrogen fixation in oxygen-deficient soils. [33]

Cyanobacteria, commonly known as blue-green algae, inhabit nearly all illuminated environments on Earth and play key roles in the carbon and nitrogen cycle of the biosphere. In general, cyanobacteria can use various inorganic and organic sources of combined nitrogen, such as nitrate, nitrite, ammonium, urea, or some amino acids. Several cyanobacteria strains are also capable of diazotrophic growth, an ability that may have been present in their last common ancestor in the Archean eon. [34] Nitrogen fixation not only naturally occurs in soils but also aquatic systems, including both freshwater and marine. Nitrogen fixation by cyanobacteria in coral reefs can fix twice as much nitrogen as on land—around 660 kg/ha/year. The colonial marine cyanobacterium Trichodesmium is thought to fix nitrogen on such a scale that it accounts for almost half of the nitrogen fixation in marine systems globally. [35]

Marine surface lichens and non-photosynthetic bacteria belonging in Proteobacteria and Planctomycetes fixate significant atmospheric nitrogen. [36]

Species of nitrogen fixing cyanobacteria in fresh waters include: Aphanizomenon and Dolichospermum (previously Anabaena). [37] Such species have specialized cells called heterocytes, in which nitrogen fixation occurs via the nitrogenase enzyme. [38] [39]

Root nodule symbioses Edit

Legume family Edit

Plants that contribute to nitrogen fixation include those of the legume family—Fabaceae— with taxa such as kudzu, clover, soybean, alfalfa, lupin, peanut and rooibos. They contain symbiotic rhizobia bacteria within nodules in their root systems, producing nitrogen compounds that help the plant to grow and compete with other plants. [40] When the plant dies, the fixed nitrogen is released, making it available to other plants this helps to fertilize the soil. [1] [41] The great majority of legumes have this association, but a few genera (e.g., Styphnolobium) do not. In many traditional farming practices, fields are rotated through various types of crops, which usually include one consisting mainly or entirely of clover. [ citation needed ]

Fixation efficiency in soil is dependent on many factors, including the legume and air and soil conditions. For example, nitrogen fixation by red clover can range from 50 to 200 lb./acre. [42]

Non-leguminous Edit

Other nitrogen fixing families include:

  • Some cycads. [citation needed]
  • Parasponia, a tropical genus in the family Cannabaceae, which are able to interact with rhizobia and form nitrogen-fixing nodules [43] such as alder and bayberry can form nitrogen-fixing nodules, thanks to a symbiotic association with Frankia bacteria. These plants belong to 25 genera [44] distributed across eight families.

The ability to fix nitrogen is present in other families that belong to the orders Cucurbitales, Fagales and Rosales, which together with the Fabales form a clade of eurosids. The ability to fix nitrogen is not universally present in these families. For example, of 122 Rosaceae genera, only four fix nitrogen. Fabales were the first lineage to branch off this nitrogen-fixing clade thus, the ability to fix nitrogen may be plesiomorphic and subsequently lost in most descendants of the original nitrogen-fixing plant however, it may be that the basic genetic and physiological requirements were present in an incipient state in the most recent common ancestors of all these plants, but only evolved to full function in some of them.

Several nitrogen-fixing symbiotic associations involve cyanobacteria (such as Nostoc):

Endosymbiosis in diatoms Edit

Rhopalodia gibba, a diatom alga, is a eukaryote with cyanobacterial N
2 -fixing endosymbiont organelles. The spheroid bodies reside in the cytoplasm of the diatoms and are inseparable from their hosts. [46] [47]

Eukaryotic Nitrogenase Engineering Edit

Some scientists are working towards introducing the genes responsible for nitrogen fixation directly into plant DNA. As all known examples of nitrogen fixation takes place in prokaryotes, transferring the functionality to eukaryotes such as plant is a challenge one team is using yeast as their eukaryotic test organism. A major problem to overcome is the oxygen-sensitivity of the produced enzymes, as well as the energy requirements. Having the process taking place inside of mitocondria or chloroplasts is being considered. [48]

The possibility that atmospheric nitrogen reacts with certain chemicals was first observed by Desfosses in 1828. He observed that mixtures of alkali metal oxides and carbon react at high temperatures with nitrogen. With the use of barium carbonate as starting material, the first commercial process became available in the 1860s, developed by Margueritte and Sourdeval. The resulting barium cyanide reacts with steam, yielding ammonia. A method for nitrogen fixation was first described by Henry Cavendish in 1784 using electric arcs reacting nitrogen and oxygen in air. This method was implemented in the Birkeland–Eyde process. [49] The fixation of nitrogen by lightning is very similar natural occurring process.

Frank-Caro process Edit

In 1898 Frank and Caro developed a way to fix nitrogen in the form of calcium cyanamide. The Frank-Caro and Ostwald processes dominated industrial fixation until the discovery of the Haber process in 1909. [50] [51]

Haber process Edit

The most common ammonia production method is the Haber process. The Haber-Bosch nitrogen reduction process for industrial fertilizer production revolutionized modern day technology. [52] Fertilizer production is now the largest source of human-produced fixed nitrogen in the terrestrial ecosystem. Ammonia is a required precursor to fertilizers, explosives, and other products. The Haber process requires high pressures (around 200 atm) and high temperatures (at least 400 °C), which are routine conditions for industrial catalysis. This process uses natural gas as a hydrogen source and air as a nitrogen source. The ammonia byproduct has resulted in an intensification of nitrogen fertilizer globally [53] and is accredited with supporting the expansion of human population from roughly 2 billion in the early 20th century to roughly 7 billion people presently. [54]

Much research has been conducted on the discovery of catalysts for nitrogen fixation, often with the goal of reducing energy requirements. However, such research has thus far failed to approach the efficiency and ease of the Haber process. Many compounds react with atmospheric nitrogen to give dinitrogen complexes. The first dinitrogen complex to be reported was Ru(NH 3)5(N2) 2+ .

Homogeneous catalysis Edit

Much research has been conducted on the discovery of catalysts for nitrogen fixation, often with the goal of reducing energy requirements. However, such research has thus far failed to approach the efficiency and ease of the Haber process. Many compounds react with atmospheric nitrogen to give dinitrogen complexes. The first dinitrogen complex to be reported was Ru(NH
3 )
5 ( N
2 ) 2+ . [55] Some soluble complexes do catalyze nitrogen fixation. [56]

Nitrogen can be fixed by lightning that converts nitrogen gas ( N
2 ) and oxygen gas ( O
2 ) present in the atmosphere into NO
x (nitrogen oxides). NO
x may react with water to make nitrous acid or nitric acid, which seeps into the soil, where it makes nitrate, which is of use to plants. Nitrogen in the atmosphere is highly stable and nonreactive due to the triple bond between atoms in the N
2 molecule. [57] Lightning produces enough energy and heat to break this bond [57] allowing nitrogen atoms to react with oxygen, forming NO
x . These compounds cannot be used by plants, but as this molecule cools, it reacts with oxygen to form NO
2 . [58] This molecule in turn reacts with water to produce HNO
3 (nitric acid), or its ion NO −
3 (nitrate), which is usable by plants. [59] [57]

Fixing a Nitrogen Deficiency in the Soil

There are two routes to go when fixing a nitrogen deficiency in the soil, either organic or non-organic.


To correct a nitrogen deficiency using organic methods requires time, but will result in a more even distribution of the added nitrogen over time. Some organic methods of adding nitrogen to the soil include:


Nitrogen as a plant fertilizer is common when purchasing chemical fertilizers. When looking to specifically add nitrogen to your garden, choose a fertilizer that has a high first number in the NPK ratio. The NPK ratio will look something like 10-10-10 and the first number tells you the amount of nitrogen. Using a nitrogen fertilizer to fix a nitrogen deficiency in the soil will give a big, fast boost of nitrogen to the soil, but will fade quickly.

Keeping the nitrogen-fixation dream alive

The conversion of inert N2 gas to a metabolically tractable form, such as ammonia, is called nitrogen fixation. In biology, nitrogen fixation is a highly oxygen-sensitive process restricted to a select group of diverse microorganisms, often collectively referred to as diazotrophs, or “nitrogen eaters.” The sparse availability of fixed nitrogen, also known as fertilizer, has historically limited worldwide food production (1 ⇓ ⇓ –4). Since about 1920, the situation has been significantly ameliorated by application of industrially produced fertilizer. Indeed, the Haber–Bosch process for industrial fertilizer production has been touted as the technological advance that has had the most impact on the modern world, driving the green revolution of the last century and fueling unprecedented population growth (1). However, the practice of applying industrially produced fertilizer to augment agricultural yield has also proven to incur severe economic, agronomic, and environmental penalties. Among these “penalites” are consumption of nonrenewable fossil fuels, prodigious production of greenhouse gases, spoiling of watersheds as a consequence of fertilizer run-off, costs associated with fertilizer distribution and application and, of course, socio-political issues associated with unbridled population growth (1 ⇓ ⇓ –4).

Emergence of the recombinant DNA era in the mid-1970s led to the ambitious goal of endowing higher plants, in particular cereal plants, with the capacity to produce their own nitrogenous fertilizers by transfer of the microbial genetic determinants to plants (5). This effort was anticipated to result in sustainable agricultural practices that could reduce the demand for production of industrial fertilizers and avoid the costs and adverse effects associated with their transportation and application. Some 40 y later, this dream remains unrealized and can be reasonably considered as biotechnology’s most significant failure. Now, with a complete understanding of the fundamental genetic determinants necessary to sustain nitrogen fixation in microbes, and at least a rudimentary understanding of the mechanistic features of how nitrogen-fixation systems are assembled and operate, together with advances in organellar targeting and the advent of synthetic biology, there has been a rejuvenated interest in revisiting the nitrogen-fixation challenge (6). Yang et al. (7) now report in PNAS the remarkable observation that certain electron-transport chains of plant origin can be recruited to provide the reducing equivalents necessary to drive nitrogen fixation. This achievement represents an incremental, although profoundly important, milestone toward realizing the goal of endowing plants with the capacity for self-fertilization.

Biological nitrogen fixation is exclusively catalyzed by a complex and extremely oxygen-sensitive metalloenzyme, called nitrogenase. Although a diverse set of microorganisms can perform nitrogen fixation, they all produce nitrogenases, which share common features with respect to catalytic mechanism and assembly of the metal-containing cofactors critical to their functions (8). Nitrogenases from all organisms described so far comprise two catalytic partners. One of those partners, designated dinitrogenase reductase, serves as a nucleotide-dependent agent of electron delivery to the other partner, dinitrogenase (9) (Fig. 1). After dinitrogenase accumulates a sufficient number of electrons from the reductase, it binds and reduces N2 to form ammonia (10) (Fig. 1). The metabolic burden of nitrogen fixation can be appreciated from the perspective that eight electrons and 16 ATP are consumed for the reduction of each N2. In addition to the nitrogenase catalytic components, there are proteins required for assembly and insertion of the metal clusters necessary to activate the catalytic unit (11). Furthermore, there are proteins required to couple cellular metabolism to nitrogen-fixation–specific electron transfer (7). Thus, there are at least two fundamental aspects that must be achieved before the production of nitrogen-fixing cereals using a direct gene-transfer approach can be considered a tractable goal. It must be demonstrated that a minimum set of genetic determinants for nitrogen fixation can be transferred to a host eukaryote and produce active nitrogenase components. Also, it must be shown that the eukaryotic host can provide the reducing power and energy necessary to sustain nitrogenase catalysis. Although neither of these objectives has been fully realized, there is now compelling proof-of-principle that both objectives can and will be achieved.

Simplified schematic of nitrogen fixation. (A, Upper) Minimal set of genes that can sustain nitrogen fixation in E. coli (9). (Lower) Eukaryotic genes that can or might functionally replace bacterial counterparts. Color schemes correspond to functional properties of the gene products shown in B. (B) Functional features of gene products shown in A and described in the text. Genes and corresponding functions are indicated by matched colors. Green arrows indicate functional replacement by eukaryotic proteins. For the bacterial system, the iron and sulfur needed to assemble all nitrogenase cofactors are supplied by the product of NifU and NifS. Cofactors are shown as ball-and-stick models: yellow (sulfur), orange (iron), black (carbon), red (oxygen). SAM 1 indicates S-adenosyl methionine. Participants of microbial origin expected to be irreplaceable are indicated by thick borders.

In recent studies it has been shown that an active dinitrogenase reducatase can be produced in eukaryotes, provided it is targeted to either mitochondria in yeast or plastids in tobacco (12, 13). These achievements are noteworthy for two reasons. First, although dinitrogenase reductase is structurally much simpler than dinitrogenase, it is much more sensitive to inactivation by oxygen. It is therefore anticipated that an active dinitrogenase can also be produced in such experimental systems if the appropriate assembly requirements are satisfied. It also appears that the much anticipated “oxygen problem” might be circumvented by mitochondrial targeting or, perhaps, plastid-targeting strategies combined with controlled conditions for expression, as suggested by Yang et al. (7). Second, dinitrogenase reductase contains an iron-sulfur cofactor that is essential for its activity (10). In nitrogen-fixing bacteria this cofactor is assembled by the products of two nitrogen-fixation–specific genes, designated nifU, encoding a cofactor assembly scaffold, and nifS, encoding a cysteine desulfurase (14). However, neither NifU nor NifS is necessary for the production of an active nitrogenase reductase in either yeast or tobacco (12, 13), indicating that counterparts involved in assembly of iron-sulfur cofactors in eukaryotic organisms—for example Isu and Nfs1 (15)—can substitute for the nitrogen-fixation–specific assembly components. It should be noted that NifU and NifS also supply the iron-sulfur building blocks necessary for formation of the complex cofactors associated with dinitrogenase (Fig. 1).

Yang et al. (7) have now addressed a similar functional replacement question but used a reverse approach: namely, they asked if plant-specific electron-transfer chains could replace the normal bacterial components. To execute these experiments, minimal gene modules were constructed that comprise the essential features necessary for nitrogen fixation to occur in the experimental microbe Escherichia coli. These include: (i) a module encoding the nitrogenase catalytic proteins, (ii) a module encoding proteins necessary for metal-containing cofactor assembly, and (iii) a module encoding proteins required to provide the reducing equivalents necessary for catalysis. Although there are a variety of different electron-transfer components that can support nitrogen fixation in microbes, the minimal electron-transport module selected for these experiments consists of two protein partners, NifJ and NifF. NifJ is a pyruvate flavodoxin oxidoreductase and NifF is a flavodoxin. NifJ couples the oxidation of pyruvate to the reduction of NifF, which, in turn, serves as a direct electron donor to dinitrogenase reductase. Yang et al. substituted NifF by various electron carriers of plastid origin, known as ferredoxins, and obtained functional hybrid electron-transfer modules (7). In this case the functional unit required that NifJ remain intact thus, a hybrid module consisting of a bacterial protein and a plastid protein is effective in coupling cellular metabolism to nitrogen fixation. It was next shown that both NifJ and NifF could be functionally replaced by intact plant plastid electron-transport chains comprised of various ferredoxin–NADPH oxidoreductases and their cognate ferredoxins. These results demonstrate that plant plastids have the capacity to couple cellular metabolism to nitrogen fixation without the need for participation of any microbial electron-transfer partner. In contrast to the situation with substitution by plastid electron-transfer modules, mitochondrial ferredoxins cannot functionally replace NifF, nor can a complete mitochondrial electron-transfer unit replace the intact NifJ–NifF module. However, a hybrid electron-transfer module consisting of a mitochondrial adrenodoxin oxidoreductase and plant-like ferredoxins could be used to replace the NifJ–NifF module, indicating that plant mitochondria might be engineered to have the capacity to support nitrogen fixation in the absence of bacterial electron-transfer components.

In the simplest model system developed so far, there are 10 proteins required to sustain nitrogen fixation in E. coli (9) (Fig. 1). With respect to the goal of transferring that capacity to plant cells, it is now established that proteins necessary for coupling cell metabolism to nitrogen fixation (NifJ and NifF) and proteins required for mobilizing iron and sulfur for metal cofactor assembly (NifU and NifS) can be supplied by the plant host. Among the 10 proteins required in the minimal microbial system is NifV, which catalyzes formation of homocitrate, an organic constituent of the metal-containing cofactor that provides the nitrogenase active site (16). Because homocitrate is a metabolite already produced by some eukaryotes (17), it might be possible to separately engineer production of homocitrate in development of the first-generation nitrogen-fixing eukaryote. Thus, in theory expression of only five microbial proteins could be required to develop a first-generation nitrogen-fixing plant (Fig. 1). In the minimal set of genes required for microbial nitrogen fixation are encoded four proteins associated with formation of the nitrogenase catalyic partners. Also included is nifB, which is an S-adenosylmethionine–dependent enzyme (11) that provides the metal-sulfur core, called NifB-co (18) (Fig. 1) of the nitrogenase active site. It seems highly unlikely that replacement of either of the catalytic components, or NifB, by proteins of strictly plant origin can ever be accomplished. Nevertheless, the current simplification of the nitrogen-fixation challenge is critically important because it lowers the number of players that must be mobilized into plant organelles and it will be easier to assess the particular function that is missing or limiting in efforts to produce plant cells that have the capacity for nitrogen fixation. In aggregate, recent advances have provided optimism that it will be possible to develop a first-generation nitrogen-fixing plant using a model system. However, the development of a truly robust system that provides significant economic and agronomic benefit will probably require a reverse of the reductionist approach used so far, and this could take yet another several decades.


Assessment of Nif Components for Polyprotein Assembly.

First, to utilize the polyprotein-based strategy, it is necessary to assess the expression levels of each component to determine which proteins are suitable for grouping together in terms of expression stoichiometry. Second, since both N-terminal and C-terminal tails can remain after TEVp cleavage, depending on positioning within the precursor polyprotein (Fig. 1A), it is necessary to determine the tailing tolerance of each gene product to design the arrangement of the coding sequences within the giant genes. Expression levels of each nif gene in their native operon locations were quantitated using in-frame translational fusions under steady-state diazotrophic conditions (SI Appendix, Fig. S1 and Table S1). Although this assay does not take into account the stability of the native nif-encoded proteins, we observed that the ratio of nifH to nifDK expression was 2:1 as demonstrated for their respective accumulated protein products in Azotobacter vinelandii (22). These relative expression levels suggest polyprotein designs in which the coding sequences of NifHDKJ, NifENBY, NifUSV, and NifFM can be grouped into giant genes that express cleavable polyproteins. Since the presence of tails after TEVp cleavage may introduce additional constraints on giant gene design, we assessed the tolerance of each Nif protein to the presence of a C-terminal ENLYFQ tail by assaying the nitrogenase activities of constructs in which the coding sequence for this tail was added to individual genes in the 18-gene reconstituted Nif system (Fig. 1B and SI Appendix, Table S2). This revealed that NifK cannot tolerate the tail and therefore can be located only at the C terminus of a polyprotein. Most of the other components were tailing-tolerant although this reduced the activity of NifB by about 30%.

To minimize the BNF system and simplify the reassignment of genes encoding polyproteins, we omitted nifT, nifX, nifW, and nifZ, which are not essential for BNF in E. coli, as single mutations in these genes do not influence nitrogenase activity (SI Appendix, Fig. S2B). Similarly, nifQ was also omitted because the function of its gene product can be recovered in the presence of a high concentration of molybdenum (SI Appendix, Fig. S2C), as observed previously (23). Guided by the expression groupings defined above and the tailing tolerance of protein components, giant genes were assembled to encode polyproteins with coding sequences flanked by TEVp recognition sites (Fig. 1B). (These giant genes are annotated from here on as nifJǒHǒDǒK, nifEǒNǒBǒY, nifUǒSǒV, and nifFǒM, where ǒ indicates the presence of a TEVp-processing site.)

Activity-Based Test and Regroup Cycles.

The functionality of our first-generation polyproteins, both before and after TEVp cleavage, was determined by measuring nitrogenase activities obtained from each giant gene when complemented with the remainder of the nif genes, respectively. Cleavage was achieved by introducing a cassette for expressing TEVp under the control of the Ptac promoter after induction with isopropyl β- d -1-thiogalactopyranoside (IPTG). Induction of TEVp expression did not influence the functionality of native Nif proteins. When assayed for acetylene reduction, the giant nifJǒHǒDǒK gene, expressed from the nifH promoter, resulted in only 5% activity after TEVp induction (SI Appendix, Fig. S3A). The nifUǒSǒV and nifEǒNǒBǒY genes also exhibited low activities of 22 and 19%, respectively (Fig. 2B and SI Appendix, Fig. S3C), in contrast to the nifFǒM gene, which showed almost full activity (99%) after TEVp induction (Fig. 2B).

Assessment of giant genes for complementation of nitrogenase activity and cleavage of their encoded polyproteins. In all cases, giant genes were expressed from native nif promoters specific to the first gene located in their coding sequences (Materials and Methods). Giant genes taken forward for assembly of the complete polyprotein-based system are indicated in red with an additional asterisk. (A) Example of a dataset for the nifHDK group. The gene arrangement for each giant gene is displayed as colored arrows with letters representing the corresponding nif gene product. TEVp sites are shown as diamonds, and linkers within fusion proteins are indicated by wavy ribbons. The acetylene reduction assay was used to measure complementation by each giant gene of the remaining nif genes in the operon-based system either in the absence of the TEVp-coding sequence (blue bars) or when TEVp expression was induced with 20 μM of IPTG (green bars). Acetylene reduction activities by the reconstituted operon-based system in E. coli were assigned as 100% (specific activities in the absence and presence of TEV were 30.4 ± 2.6 and 29.2 ± 1.7 nmol C2H4/min/mg total protein, respectively). Error bars indicate the SD observed from at least two biological replicates. Samples were immediately collected after the acetylene reduction assay for Western blotting (antibodies used are listed in Materials and Methods. Complete gels of the Western blots are provided in SI Appendix, Fig. S9). “EV” represents empty vector, used as a negative control. Image J software was used for quantification of NifH protein, and relative expression levels are shown in red as a percentage (in parentheses). (B) Summary data for the nifENB, nifUSV, nifJVWZ, nifUSZ, and nifFMY groups. The gene arrangement is shown in short format with rows b1, c1, d1, e1, and f1 representing the original operon-based system in E. coli. Commas indicate separate operons, “ǒ” indicates a TEVp site, and “∼” indicates a fusion protein. “†” indicates SD values lower than 0.5. Complete datasets for these groupings including the associated Western blots are shown in SI Appendix, Fig. S4.

The low activities of proteins following cleavage from polyproteins prompted us to test multiple gene combinations in a series of regrouping and retesting cycles. We observed that coexpression of NifJ with NifHDK led to decreased protein levels of NifH, NifD, and NifK compared with proteins expressed from the native nifHDK operon, although the cleavage products appeared similar (SI Appendix, Fig. S3B). Consequently, when nifJ was excluded from the giant gene, the resultant nifHǒDǒK assembly enabled 75% nitrogenase activity after TEVp cleavage of its polyprotein product, with a similar NifD:NifK protein stoichiometry to that expressed from the native genes (Fig. 2A). As NifB showed weak tolerance (71%) to the C-terminal ENLYFQ-tail (SI Appendix, Table S2), we removed nifY from nifEǒNǒBǒY, and the resultant nifEǒNǒB gene restored 72% of nitrogenase activity, following TEVp induction (Fig. 2B). When the NifY-coding sequence was reassigned to nifFǒM, cleavage of the larger polyprotein encoded by nifFǒMǒY resulted in 89% of the nitrogenase activity exhibited by the native components, and increased levels of NifY were observed (Fig. 2B and SI Appendix, Fig. S4E). For the nifUSV group, posttranslational splicing of the NifUǒSǒV polyprotein led to decreased levels of NifU and nearly undetectable amounts of NifS, but the level of NifV did not apparently change (compare lanes c1 and c2 in SI Appendix, Fig. S4B). In an attempt to improve this, we removed the nifV sequence from the giant gene, but retained NifUS coexpression with nifUǒS. Surprisingly, this restored native levels of NifU and NifS after cleavage and recovered nitrogenase activity to 82% (compare lanes c2 and c5 in SI Appendix, Fig. S4B). To assign nifV to another giant gene, we regrouped it with nifJ to form nifJǒV. Interestingly, in this case the polyprotein product was active even before protease cleavage, resulting in 65% nitrogenase activity, which increased to 95% after splicing with TEVp. Although native levels of NifV were released under these conditions, the amount of NifJ decreased (compare lanes d1 and d2 in SI Appendix, Fig. S4C).

To further optimize activity, we carried out additional regrouping of genes encoding polyproteins and also tested the incorporation of fused genes as a means to simplify the ensembles. Since splicing of NifHǒDǒK by TEVp resulted in a decreased amount of NifH (66%), but in wild-type levels of NifD and NifK (compare lanes a1 and a2 in Fig. 2A), we attempted to restore the optimal 2:1 ratio of NifH:NifDK (22) through assembly of a giant gene (nifHǒHǒDǒK) expressing two copies of nifH. Although this ensemble slightly increased the level of NifH after cleavage (70%), it did not result in increased nitrogenase activity (compare lanes a2 and a4 in Fig. 2A). To further attempt optimization of NifHDK levels, we also incorporated fusion proteins with different linkers guided by previous studies and natural existing examples (24 ⇓ ⇓ –27). Fused NifD∼K proteins showed broad tolerance to different lengths of GGGGS linkers, with a maximum activity of 91% when 5× GGGGS linkers were added (SI Appendix, Fig. S5A). We also found that two copies of the NifH protein could be functionally fused with an ArsA linker retaining 89% nitrogenase activity (SI Appendix, Fig. S5B). However, integrating NifD∼K or NifH∼H fusions into two further ensembles (nifHǒD∼K and nifH∼HǒDǒK) did not result in higher activities than the original nifHǒDǒK gene (compare lane a2 with lanes a3 and a5 in Fig. 2A).

For the nifENB group, functional fusions of NifE∼N and NifN∼B were obtained using 5× and 3×GGGGS linkers, which exhibited activities of 91 and 115%, respectively (SI Appendix, Fig. S5 C and D). We replaced the corresponding parts in the nifEǒNǒB gene to generate three more assemblies (nifE∼NǒB, nifEǒN∼B, and nifE∼N∼B). The incorporation of either the NifE∼N or NifN∼B fusion resulted in higher nitrogenase activities (76 and 89%, respectively) compared with the nifEǒNǒB gene (compare lanes b3 and b4 with b2 in SI Appendix, Fig. S4A). However, when all three genes were fused to express the NifE∼N∼B protein, only 50% nitrogenase activity was obtained. This decrease may reflect the presence of truncated NifE∼N translation products expressed from nifE∼N∼B (see lane b5 in SI Appendix, Fig. S4A). We also attempted fusion of NifU with NifS and obtained 50% nitrogenase activity with a 5× GGGGS linker when expressed in the native operon arrangement nifU∼SVWZM (SI Appendix, Fig. S5E). However, this fusion protein had lower activity than that obtained after cleavage of the NifUǒS polyprotein (compare lanes c3 and c5 in SI Appendix, Fig. S4B and lanes e2 and e3 in SI Appendix, Fig. S4D).

Assembly and Characterization of Complete Polyprotein-Based Nitrogenase Systems.

To combine polyproteins into a functional Nif system, we sequentially replaced the native gene parts with giant gene assemblies (Fig. 3A). Sequential combination of nifHǒDǒK with nifFǒMǒY and nifEǒN∼B (thus reducing the number of genes from 16 to 9) resulted in relatively small decreases in nitrogenase activity as measured both by acetylene reduction and 15 N assimilation (Fig. 3A, rows I–IV). However, replacement of the native nifUSVWZ genes with nifUǒSǒV (thus reducing the number of genes to 5) resulted in a dramatic decrease in activity (10% of the native system) when acetylene reduction was measured (compare rows IV and V in Fig. 3A). The greater influence on acetylene reduction is probably the result of a kinetic effect as the 15 N assay was conducted over a longer time window. Nitrogenase activity improved when nifV was switched from nifUǒSǒV to nifJǒV in the five-gene system (Fig. 3A, compare rows V and VI) as anticipated from the analysis of single polyproteins (Fig. 2B). Nevertheless, the decreased activity observed in the absence of nifW and nifZ prompted us to reconsider their involvement. Although the nifWZ gene products do not apparently have an impact on the activity of our reconstituted system (SI Appendix, Fig. S2B), previous studies suggest that they are required for full activity of the MoFe protein (28, 29). Bearing in mind the native location and expression levels of nifW and nifZ, we assembled additional giant genes designed to express their gene products as polyproteins with NifJ and NifV (nifJǒVǒZ, nifJǒVǒW, and nifJǒVǒWǒZ). When assayed to complement the native genes, the highest activity was obtained with the polyprotein expressing NifJVW (98%), and no benefit was obtained by incorporating NifZ (SI Appendix, Fig. S4C, compare lanes d3, d4, and d5). These activities were mirrored when these giant genes were complemented with the other four genes encoding polyproteins, with nifJǒVǒW again giving the highest level of activity (51% for acetylene reduction compare arrangements VI, VII, VIII, and IX in Fig. 3A).

Assembly and characterization of the polyprotein-based nitrogenase system. (A) Schematic diagram showing the process of assembly by replacing native genes with regrouped giant genes. Numbers in parentheses on the left represent gene numbers (including giant genes and native genes) for each construct. Each ensemble was analyzed by both acetylene reduction and 15 N assimilation to measure nitrogen fixation activity. The activities exhibited by the reconstituted operon-based system in E. coli [the pKU7017 plasmid is assembled with seven operon-based biobricks (5)] were assigned as 100% (29.1 ± 0.8 nmol C2H4/min/mg total protein for acetylene reduction assay and 1,172 ± 75 δ 15 N/ 14 N‰ for the 15 N assimilation assay) (row I). Error bars indicate the SD observed from at least two biological replicates. (B) Mass spectrometry analysis of protein levels from samples taken immediately after the acetylene reduction assays. The yellow bars represent protein samples from a nitrogen-fixing culture of K. oxytoca, grown under the same conditions as the E. coli cultures the blue bars represent protein samples from the reconstituted operon-based nif system in E. coli (construct I in A), and the green bars represent protein samples from the polyprotein-based nif system (construct VIII in A) the GAPDH protein encoded by the E. coli gapA gene (EcgapA) was assigned as the internal reference. Asterisks mark proteins that could not be assigned (for NifK, chemical synthesis of the internal standard peptides failed, and, for NifW, no peptide with a detectable signal was identified). Error bars indicate the SD observed from three biological replicates. (C) Diazotrophic growth promoted by polyprotein-based nitrogenase systems in E. coli in the presence of 20 μM of IPTG. Roman numerals represent the corresponding assemblies in A. EV represents empty vector, used as a negative control. Control plates with strains grown in the absence of IPTG are shown in SI Appendix, Fig. S7.

Quantitative analysis of protein levels by Selected Reaction Monitoring (SRM) mass spectrometry revealed that, overall, the stoichiometry of most components from the polyprotein-based system matched remarkably well with the respective levels from the reconstituted operon-based system in E. coli and the native system in the original K. oxytoca host (Fig. 3B compare green, blue, and yellow bars, respectively). This was particularly true for the NifHDK and NifENB proteins, where stoichiometry is important for nitrogenase biosynthesis and activity [the level of NifK was determined by quantification of Western blots (SI Appendix, Fig. S6)]. Unexpectedly, NifU and NifS levels in the polyprotein system were more similar to those in the native K. oxytoca host, compared with the original recombinant system in E. coli (Fig. 3B and SI Appendix, Fig. S6). Although expression of NifU and NifS as a single polyprotein limits complementation of a nifUS deletion (Fig. 2B), only a small benefit was achieved when nifU and nifS were included as separate genes, with the other four polyproteins (compare rows VIII and X in Fig. 3A). Although NifJ and NifV were located on the same polyprotein, the level of NifV was fivefold higher than that of NifJ, which is similar to the ratio obtained from the native system. This suggests that NifJ may have a relatively short half-life, which is not reflected when expression is measured at the transcriptional level (SI Appendix, Table S1). However, the rearrangement of nifF, nifM, and nifY into a giant gene (nifFǒMǒY) resulted in increased levels of all three protein products, as anticipated from their inclusion in the same polyprotein (Fig. 3B). This may have consequences for the level of NifH, which, unexpectedly, did not decrease in the polyprotein-based system in contrast to the data obtained in Fig. 2A. The increased level of NifM resulting from posttranslational splicing of the NifFMY polyprotein may be responsible for the increased NifH accumulation since NifM is required for the maturation of NifH (17).

Since the combined five-gene (Fig. 3A, row VIII) and six-gene (Fig. 3A, row X) polyprotein systems exhibited 72 and 75% 15 N assimilation activity, respectively, we anticipated that these combinations of giant genes could support diazotrophic growth by E. coli. As in the case of the initial single-gene system (Fig. 3A, row I), the five-gene and six-gene arrangements (assemblies VIII and X, respectively) enabled E. coli to grow on solid media with dinitrogen as the sole nitrogen source, when the expression of TEVp was induced by IPTG. In contrast, assemblies IX and XI, which exhibited lower nitrogenase activates, grew less well under these conditions (Fig. 3C). Control experiments in the absence of IPTG, which results in basal levels of TEVp, resulted in poor growth of assemblies VIII and X as anticipated (SI Appendix, Fig. S7).


The tubers are highly palatable with culinary characteristics of a potato, although the flavor can be somewhat nuttier than a potato and the texture can be finer. [5] Studies in rats suggest that raw tubers should not be consumed. They contain harmful protease inhibitors that are denatured by cooking. [6] These tubers contain roughly three times the protein content of a potato (16.5% by dry weight), and the amino acid balance is good with the exception of cysteine and methionine. [7] Apios americana tubers were found to have a protein concentration of 15–30 mg/g (0.24–0.48 oz/lb). [8] This was similar to that of other species in the Apios genus, A. carnea and A. fortunei. [8] However, A. americana had larger levels of genistein than the other two species. [8] The fatty acid content of tubers is approximately 4.2% to 4.6%, with linoleic fatty acids predominating. [9] Thirty-six percent of the fresh weight of a tuber is carbohydrate (primarily starch). [10] The tubers are also an excellent source of calcium and iron. [10] Calcium content is 10-fold greater than a potato and iron is 2-fold greater than a potato, although vitamin C was considerably less than a potato. [10] The tuber and the flower also contain mono and oligosaccharides. [11] The tuber has more monosaccharides and oligosaccharides than the soybean, potato, and sweet potato. [11]

In addition, the tubers appear to have numerous health-promoting factors. Hypertensive rats that were fed powdered tubers as 5% of their total diet had a 10% decrease in blood pressure and also a reduction in cholesterol and triglycerides. [12] It has been shown that the tubers contain genistein and other isoflavones that have various health benefits, including an anti-carcinogenic function against colon, prostate, and breast cancer. [13] [14] Genistein-7-O-gentiobioside is a novel isoflavone that is found in the American groundnut. [15] Extract from the American groundnut was shown to drive the anti-oxidative pathway in cells although it did not have anti-oxidative activity itself. [15] Human breast carcinoma MCF-7 cells were pretreated with the extract of A. americana for 24 hours. [15] Subsequent analysis showed an increase in expression of heme oxygenase-1, a protein induced during oxidative stress. [15] The American groundnut, like soybean, is a great source of isoflavone. [15]

Furthermore, a study on A. americana and its flower shows that the flower of the particular plant is not toxic to mice. [16] Consumption of the flower was shown to lower plasma glucose levels in diabetic mice. [16] The flower was shown to have an inhibitory activity on maltose and an anti-hyperglycemic effect in mice, suggesting that not only is it a viable and novel food source for the general population, but also in the prevention of diabetes.

The only place in the world today where American groundnuts are commercially farmed in any significant quantities is in Japan. [ citation needed ] Before the American groundnut was introduced to Japan, the people on the main island of Honshu and the northern island of Hokkaido were already familiar with a native, wild plant called hodoimo (Apios fortunei), which was occasionally eaten as an emergency food. [10] It is believed that sometime during the Meiji period (1868-1912), American groundnut was accidentally or deliberately brought to Japan. [10] [12] [14] [17] [18] One theory is that American groundnut was accidentally brought to Japan as a stowaway weed among apple seedlings imported from North America. [12] [14] Another theory is that American groundnut may have been deliberately brought to Japan in the middle of the Meiji period as an ornamental flower. [10] [17]

It has become a culinary specialty of the Aomori Prefecture, where American groundnut agriculture is centered. It has been eaten there for more than one hundred years. [19] [20] Although American groundnut agriculture is primarily identified with agriculture in the Aomori prefecture, it is grown in the nearby prefectures of Akita and Miyagi as well. [18] In addition, it is known to be grown in the southern part of Honshu in the Tottori prefecture. Radioactive testing records following the Fukushima nuclear disaster record cesium testing of American groundnut agricultural products in the central prefecture of Tochigi. [18] [21]

An important part of the spread and popularization of American groundnut consumption in Japan has been the efforts of Dr. Kiyochika Hoshikawa to promote the cultivation of this crop in Japan, and the flurry of scientific articles on the health benefits of eating American groundnut tubers. [12] Japanese websites that sell American groundnut continue to emphasize its health benefits in their marketing efforts. [20] [22] There are reports of American groundnut cultivation in South Korea as well, where it is grown for its nutritional benefits. [23]

The tubers and seeds can both be cooked and eaten. [24]

By Indigenous peoples of the Americas Edit

The tubers have traditionally been a staple food among most Indigenous peoples of the Americas within the natural range of the plant. [25] In 1749, the travelling Swedish botanist Peter Kalm writes, "Hopniss or Hapniss was the Indian name of a wild plant, which they ate at that time. The roots resemble potatoes, and were boiled by the Indians who ate them instead of bread." [25] Strachey in 1612 recorded observations of the Indigenous peoples found in Virginia: "In June, July, and August they feed upon roots of tockohow, berries, groundnutts, fish, and greene wheate. " [25] In Eastern Canada, the Jesuit missionary, Le Jeune, observed that the Indigenous peoples there would, "eat, besides, roots, such as the bulbs of the red lily . another that our French people call 'Rosary' because it is distinguished by tubers in the form of beads." [25] The early author Rafinesque observed that the Cree were cultivating the plant for both its tubers and seeds. [25] The author Brinton wrote in 1885 in regards to the Lenape people, "Of wild fruits and plants they consumed the esculent and nutritious tubers on the roots of the Wild Bean, Apios tuberosa. which the Indians called hobbenis. " [25] In 1910, Parker writes that the Iroquois were consuming significant quantities of groundnuts up until about thirty years before his writing. [25] The Paris Documents of 1666 record that the sixth tribe of the second division of the Iroquois were identified as, "that of the Potatoe, which they call Schoneschironon" and an illustration of tubers is found in the Paris Documents with the explanation, "This is the manner they paint the tribe of the Potatoe." [25] The author Gilmore records the use of groundnuts by the Caddoan and Siouan tribes of the Missouri river region, and the authors Prescott and Palmer record its use among the Sioux. [25] The Indigenous peoples would prepare the tubers in many different ways, such as frying them in animal fat or drying them into flour. [26] Many tribes peel them and dry them in the sun, such as the Menomini who have traditionally built scaffolds of cedar bark covered with mats to dry their tubers for winter use. [25] The Menomini are recorded as having dried the tubers in maple syrup or making a preserve of Groundnut tubers by boiling them in maple syrup. [25] The Potawatomi have traditionally boiled their tubers. [25] The traditional Meskwaki and Chippewa preparation involves peeling, parboiling, slicing, and drying the tubers. The Chippewa have historically used them as a sort of seasoning in all their foods. [25]

By Europeans Edit

The Europeans learned to use the American groundnut from the Indigenous peoples of the Americas. As a result, the American groundnut became interwoven with the history of the American colonies and Europe. The early traveler John Brereton was sustained by the "good meat" and "medicinable" qualities of American groundnut during his travels in New England in 1602. [25] In 1613, the followers of Biencourt at Port-Royal ate the tubers to help them survive in the New World. [25] The American groundnut was an important factor in the survival of the Pilgrims during the first few winters of their settlement. [27] In 1623 the Pilgrims, "having but a small quantity of corn left," were "enforced to live on groundnuts. and such other things that the country afforded. and were easily gotten. ". [25] The Pilgrims were taught to find and prepare American groundnut by the Wampanoag people. [5] The groundnut was likely eaten at the harvest festival of November 1621 that is regarded as the first Thanksgiving, although only venison was specifically named as a food item at this meal by a Pilgrim eyewitness account.

Philosopher Henry David Thoreau commented on the nutty flavor and dry texture in October 1852. [26]

It is believed that American groundnut may have been shipped to Europe as early as 1597. It was listed in 1885 as a European garden crop. [2] In 1845 it was evaluated as a possible alternative potato crop in Ireland during the Great Famine. [2] These early introductions to Europe appear to have resulted in little or no assimilation of the new food into the European diet. [2] A primary reason for this lack of assimilation was that the two-year cycle for an acceptable tuber yield did not match the cropping systems that were familiar to Europeans. [2]

American groundnut is generally considered to be an undomesticated crop. In her 1939 description of the Native American use of American groundnut, Gretchen Beardsley states that several historical sources describe the "cultivation" of American groundnut by Indigenous peoples. She dismisses the ambiguous term "cultivation" as perhaps referring to transplantation of tubers near a settlement. She quotes the historical author Waugh on this subject of cultivation: "sometimes planted in suitable locations, though they are not, strictly speaking, cultivated." [25] Subsequent authors on the American groundnut have followed Beardsley's interpretation of "cultivation" when referring to the early use by Indigenous peoples of American groundnut. However, recent evidence suggests that North American indigenous peoples likely intervened significantly as cultivators of the native plants of the region, in a manner similar to contemporary Western permaculture practices. [28] So, from a permacultural perspective, Native Americans may well have "cultivated" the groundnut.

In 1985, Dr. William J. Blackmon, Dr. Berthal D. Reynolds, and their colleagues at Louisiana State University in Baton Rouge began a program of deliberate domestication of American groundnut. Their primary goal was to develop an American groundnut that can produce a significant yield in a single season. [2] [5] [27] Early trials identified LA85-034 as a promising cultivar, with "elongate tubers of uniform, medium size with light brown skin and little extra rhizomatous material". [27] By 1988, they had collected wild seeds and tubers from 210 plants found in 19 states, although the bulk of their selections came from the state of Louisiana. [2] From these wild materials, and a small number of single crosses, they rigorously selected for plants that met their primary breeding goals of (1) larger tuber size, (2) denser tuber set, (3) single season production, and (4) productivity in untrellised cultivation. [2] The American groundnut domestication program at Louisiana State University continued in various forms until the mid-1990s. [5] Cultivars from this program can still occasionally be found available from small seed companies.

From 1985 to 1994, an Apios breeding program took place that resulted in the collection of over two hundred wild accessions. These accessions underwent hybridization and selection, and over 2,200 lines were assessed. [29] Of these lines, only 53 genotypes were kept for further analysis. Three different locations and three different growing conditions—field, pots, and grow-bags— were used. There was significant variation found among almost all of the 20 genotypes in the field growing condition. Inter-node length, plant vigor, and stem diameter during plant growth were positively correlated with the plant yield below ground. [29] There were four distinct genotypic clusters found in this collection of Apios lines. Several genotypes yielded large plants in all locations, maxing up to 1.5 kg (3.3 lb) of below ground tuber. [29] This suggests that the plant has a good ability to adapt and grow in a wide variety of locations and conditions. Furthermore, the superior germplasm identified in this project may be suitable as cultivars, and will aid in further development of Apios lines as a crop. [29]

The largest germplasm collection of Apios americana cultivars today is found at Iowa State University under the direction of Dr. Steven Cannon. [23] It is maintained there for scholarly and academic use. Research continues at Iowa State on the domestication of American groundnut. [23] [30]

Despite these efforts at domestication, the American groundnut remains largely uncultivated and underused in North America and Europe. [5] There are challenges to breeding and domesticating this plant, as well. [31] There seems to be a partial self-incompatibility with Apios breeding and manual pollinations, resulting in rare seed-sets. [31] Disadvantages in Apios as a crop are its vining habit. [32] The crop has small tuber size for most genotypes. [32] These sizes are typically smaller than 50 g (1.8 oz) however, some do average around 100 g (3.5 oz). [32] The tuber plant is difficult to harvest because of the "beads on a string" arrangement on stolons, which extend for over a meter. [32]

American groundnut fixes its own nitrogen, which could be a great advantage in comparison to other roots crops, such as potatoes, true yams, and sweet potatoes. These do not fix their own nitrogen and require large applications of nitrogen fertilizer. [33] American groundnut can be nodulated by bacterial strains that are normally found in symbiosis with soybeans or cowpeas. [33]

Research has been done on the potential of the soybean strain B. japonicum to nodulate American groundnut. [33] It was found that plants nodulated with B. japonicum yielded

30% better than unnodulated plants if no nitrogen fertilizer was used. [33] It was also determined that nodulated plants partitioned more carbon into non-edible shoots when they were given nitrogen fertilizer, whereas unnodulated plants responded to nitrogen fertilizer with greater tuber yields than nodulated plants. [33] This data suggests that nitrogen fertilization may be required to maximize tuber size and yields in A. americana. [33]

American groundnut is normally 2n=2x=22, diploid, but both diploid and triploid forms exist. [2] Only diploids are capable of producing seeds triploids will produce flowers but not seeds. [34] Thus, triploids are entirely dependent on tuber division for propagation whereas diploids can be propagated through both seeds and tubers. [34] Other than seed production, there are no easily identifiable differences between diploids and triploids. [34] Triploids are generally found in the Northern part of American groundnut's range whereas diploids predominate in the Southern part of the range. [34] Triploids have been identified in the provinces or states of New Brunswick, Quebec, Ontario, Connecticut, Vermont, Massachusetts, New York, Pennsylvania, Ohio, New Hampshire, Rhode Island, Wisconsin, and Iowa. [34] [35] [36] A few diploids have been found in the Northeastern part of the range, such as along the Black River in Central Ontario. [35] All samples tested in the Southeastern United States have been found to be diploid. [34] [35] [36]

Nitrogen fixation in nature

Nitrogen is fixed, or combined, in nature as nitric oxide by lightning and ultraviolet rays, but more significant amounts of nitrogen are fixed as ammonia, nitrites, and nitrates by soil microorganisms. More than 90 percent of all nitrogen fixation is effected by them. Two kinds of nitrogen-fixing microorganisms are recognized: free-living (nonsymbiotic) bacteria, including the cyanobacteria (or blue-green algae) Anabaena and Nostoc and genera such as Azotobacter, Beijerinckia, and Clostridium and mutualistic (symbiotic) bacteria such as Rhizobium, associated with leguminous plants, and various Azospirillum species, associated with cereal grasses.

The symbiotic nitrogen-fixing bacteria invade the root hairs of host plants, where they multiply and stimulate the formation of root nodules, enlargements of plant cells and bacteria in intimate association. Within the nodules, the bacteria convert free nitrogen to ammonia, which the host plant utilizes for its development. To ensure sufficient nodule formation and optimum growth of legumes (e.g., alfalfa, beans, clovers, peas, and soybeans), seeds are usually inoculated with commercial cultures of appropriate Rhizobium species, especially in soils poor or lacking in the required bacterium. (See also nitrogen cycle.)

How do plants get their nitrogen from the air?

Plants do not get their nitrogen directly from the air. Although nitrogen is the most abundant element in the air, every nitrogen atom in the air is triple-bonded to another nitrogen atom to form molecular nitrogen, N2. This triple bond is very strong and very hard to break (it takes energy to break chemical bonds whereas energy is only released when bonds are formed). As a result, even though nitrogen in the air is very common, it is energetically unfavorable for a plant to split the nitrogen molecule in order to get the raw atoms that it can use. The strong triple bond of N2 also makes it hard for molecular nitrogen to react with most other chemicals. This is, in fact, part of the reason there is so much nitrogen in the air to begin with. Also, the stability and symmetry of the nitrogen molecule makes it hard for different nitrogen molecules to bind to each other. This property means that molecular nitrogen can be cooled to very low temperatures before becoming liquid, leading liquid nitrogen to be a very effective cryogenic liquid.

The act of breaking apart the two atoms in a nitrogen molecule is called "nitrogen fixation". Plants get the nitrogen that they need from the soil, where it has already been fixed by bacteria and archaea. Bacteria and archaea in the soil and in the roots of some plants have the ability to convert molecular nitrogen from the air (N2) to ammonia (NH3), thereby breaking the tough triple bond of molecular nitrogen. Such organisms are called "diazotrophs". From here, various microorganisms convert ammonia to other nitrogen compounds that are easier for plants to use. In this way, plants get their nitrogen indirectly from the air via microorganisms in the soil and in certain plant roots. Note that lightning and high-energy solar radiation can also split the nitrogen molecule, and therefore also fixes the nitrogen in the air. However, the amount of nitrogen fixed by lightning and solar radiation is insignificant compared to the amount fixed by diazotrophs in the soil and in roots. In his book Nitrogen Fixation, John Postgate states,

The fixation of nitrogen – the conversion of atmospheric nitrogen into a form that plants can use – is a process fundamental to world agriculture. It comes about as a consequence of spontaneous, anthropogenic and biological activities. The existence and importance of the biological component have been recognized for more than a century, but scientific advances over the past few decades have radically altered our understanding of its nature and mechanisms.

Human alteration of the N cycle and its environmental consequences

Early in the 20 th century, a German scientist named Fritz Haber figured out how to short-circuit the nitrogen cycle by fixing nitrogen chemically at high temperatures and pressures, creating fertilizers that could be added directly to soil. This technology spread rapidly over the 20 th century, and, along with the advent of new crop varieties, the use of synthetic nitrogen fertilizers led to an enormous boom in agricultural productivity. This agricultural productivity has helped us to feed a rapidly growing world population, but the increase in nitrogen fixation has had some negative consequences as well. While the consequences are perhaps not as obvious as an increase in global temperatures (see our Data Analysis and Interpretation module) or a hole in the ozone layer (see The Practice of Science module), they are just as serious and potentially harmful for humans and other organisms.

Why? Not all of the nitrogen fertilizer applied to agricultural fields stays to nourish crops. Some is washed off of agricultural fields by rain or irrigation water, where it leaches into surface water or groundwater and can accumulate. In groundwater that is used as a drinking water source, excess nitrogen can lead to cancer in humans and respiratory distress in infants. The US Environmental Protection Agency has established a standard for nitrogen in drinking water of 10 mg per liter nitrate-N. Unfortunately, many systems (particularly in agricultural areas) already exceed this level. By comparison, nitrate levels in waters that have not been altered by human activity are rarely greater than 1 mg/L. In surface waters, added nitrogen can lead to nutrient over-enrichment, particularly in coastal waters receiving the inflow from polluted rivers. This nutrient over-enrichment, also called eutrophication, has been blamed for increased frequencies of coastal fish-kill events, increased frequencies of harmful algal blooms, and species shifts within coastal ecosystems.

Reactive nitrogen (like NO3 - and NH4 + ) present in surface waters and soils, can also enter the atmosphere as the smog-component nitric oxide (NO) which is a component of smog, and also as the greenhouse gas nitrous oxide (N2O). Eventually, this atmospheric nitrogen can be blown into nitrogen-sensitive terrestrial environments, causing long-term changes. For example, nitrogen oxides comprise a significant portion of the acidity in acid rain, which has been blamed for forest death and decline in parts of Europe and the northeastern United States. Increases in atmospheric nitrogen deposition have also been blamed for more subtle shifts in dominant species and ecosystem function in some forest and grassland ecosystems. For example, on nitrogen-poor serpentine soils of northern Californian grasslands, plant communities have historically been limited to native species that can survive without a lot of nitrogen. There is now some evidence that elevated levels of atmospheric N input from nearby industrial and agricultural development have allowed invasion of these ecosystems by non-native plants. As noted earlier, NO is also a major factor in the formation of smog, which is known to cause respiratory illnesses like asthma in both children and adults.

Currently, much research is devoted to understanding the effects of nitrogen enrichment in the air, groundwater, and surface water. Scientists are also exploring alternative agricultural practices that will sustain high productivity while decreasing the negative impacts caused by fertilizer use. These studies not only help us quantify how humans have altered the natural world, but increase our understanding of the processes involved in the nitrogen cycle as a whole.


Although the majority of the air we breathe is N2, molecular nitrogen cannot be used directly to sustain life. This module provides an overview of the nitrogen cycle, one of the major biogeochemical cycles. The five main processes in the cycle are described. The module explores human impact on the nitrogen cycle, resulting in not only increased agricultural production but also smog, acid rain, climate change, and ecosystem upsets.

Key Concepts

The nitrogen cycle is the set of biogeochemical processes by which nitrogen undergoes chemical reactions, changes form, and moves through difference reservoirs on Earth, including living organisms.

Nitrogen is required for all organisms to live and grow because it is the essential component of DNA, RNA, and protein. However, most organisms cannot use atmospheric nitrogen, the largest reservoir.

The five processes in the nitrogen cycle – fixation, uptake, mineralization, nitrification, and denitrification – are all driven by microorganisms.

Humans influence the global nitrogen cycle primarily through the use of nitrogen-based fertilizers.

Watch the video: MEPs press conference on the abusive use of Green Certificate - European Parliament (December 2022).