Nitrogen Fixation by US and Middle Eastern Chickpeas with Commercial and Wild Middle Eastern Inocula

Abi-Ghanem, Rita; Carpenter-Boggs, Lynne; Smith, Jeffrey L.; Vandemark, George J.

doi:https://doi.org/10.5402/2012/981842

International Scholarly Research Notices

On this page

Abstract Introduction Materials and Methods Results Discussion Conclusion Abbreviations References Copyright Related Articles

Research Article | Open Access

Volume 2012 | Article ID 981842 | https://doi.org/10.5402/2012/981842

Nitrogen Fixation by US and Middle Eastern Chickpeas with Commercial and Wild Middle Eastern Inocula

Rita Abi-Ghanem,¹Lynne Carpenter-Boggs,²Jeffrey L. Smith,³and George J. Vandemark⁴

Academic Editor: Y. Feng

Received16 Dec 2011

Accepted29 Jan 2012

Published29 Feb 2012

Abstract

Chickpeas (Cicer arietinum L.) are native to the Middle East (ME) and must be inoculated with symbiotic bacteria (Mesorhizobium ciceri) to fix nitrogen (N) in North American soils. Whether commercial M. ciceri strains are more or less effective than wild strains from ME soils when paired with various chickpea hosts must be elucidated. Wild N-fixing bacterial strains were isolated from ME soils, and their effectiveness was compared against commercial strains on US and ME chickpea varieties. Chickpeas were inoculated with individual strains and grown in chambers for 8 weeks. Plants received 2 mM (¹⁵NH₄)₂ SO₄ (5% atom excess) to measure N fixation by isotope dilution. Plant below- and above-ground biomass and proportion of N fixed (PNF) were determined. Commercial and wild ME strains were examined for genetic diversity by sequencing their 16 S rDNA region. The PNF was significantly influenced by inoculant strains and chickpea varieties. Among varieties, Sierra, Troy, and Almaz had the highest PNF of 86.7%, 85.3%, and 85.2%, respectively. Among strains, Jord-M1 contributed to greater PNF (84.7%) compared to Syr-M1 (81.4%). Overall, chickpea varieties had greater effect on PNF than strain selection. These findings support efforts focusing on varietal breeding and strain selection to increase agricultural N fixation.

1. Introduction

Global demand for nitrogen fertilizer is predicted to increase 1.4% annually, which includes a 23.1 million tonnes increase by 2012 [1]. In the US, N fertilizer costs approximately tripled from 2002–2011 [2]. Although prices have decreased somewhat from the 2008 peak, the loss of billions of dollars in farm profit has drawn attention to the need for alternative sources of N. The primary feedstock for synthetic nitrogen fertilizer is natural gas, both as a source of hydrogen and energy [3]. The US is estimated to have only 3.7% of the world natural gas reserves [4], which suggests that American agriculture would benefit from less expensive and more sustainable N sources.

Chickpea (Cicer arietinum L.), a drought-resistant legume from the Middle East (ME), has been cultivated for over 10,000 years [5]. Chickpeas belong to two major types, desi and kabuli. Desi varieties have a small dark seed with a rough coat while kabuli seeds are larger with lighter color and smoother coat [6]. In the US, 51,435 ha were planted with chickpeas in 2011 with a total production of 103,179 tonnes [7]. Chickpeas form root nodules that support biological N fixation (BNF) and host symbiotic N-fixing bacteria. On average the estimated amounts of N fixed by chickpeas under regular precipitation and drought stress conditions are 60 kg/ha [8] and 19–24 kg/ha [9], respectively.

There is increasing evidence that suggests that more N can be fixed by existing legume grain crops if they are inoculated more often or with more effective strains of rhizobia [10]. Significant increases in BNF can result from the use of different commercial or wild strains [11]. The efficiency of biological N fixation is also influenced by cultivars’ selection; significant differences were observed among pea cultivars for the percentage of plant N supplied by bacterial N fixation and also for the number of root nodules formed per plant in a greenhouse experiment [12]. The known N-fixing symbiont of chickpea, M. ciceri, is not native to North American soils, and the number of commercial strains is limited. In addition, there has been little work to enhance the ability of chickpeas to support N fixation [13]. Plant breeders may be able to improve chickpea varieties through the use of breeding lines that are better N-fixing symbiont hosts and could increase the BNF process.

The objectives of this study were to isolate and identify new N-fixing symbiont strains for chickpea from ME soil and to compare the BNF effectiveness of the new strains and current commercial strains when paired with US and ME varieties of chickpeas.

2. Materials and Methods

2.1. N Fixation Study

Soil was collected at 10–15 cm depth from non-inoculated chickpea fields in the following Middle Eastern cities with the help of the following collaborating entities: (1) Terbol, Lebanon (American University of Beirut (AUB)), (2) Tal Hadya, Syria (International Center for AgriculturaI Research in the Dry Areas (ICARDA)), (3) Erbid, Jordan (National Center for Agricultural Research and Technology Transfer of the Ministry of Agriculture (NCARTT)), and (4) Yozgat, Turkey (International Maize and Wheat Improvement Center (CYMMIT)). To isolate symbiotic BNF bacteria from these soils, chickpea seeds (Arman and Sierra) were sterilized with 0.5% sodium hypochlorite for three minutes and germinated for 5 days on sterile filter paper. Germinated seeds were planted in 164 mL seedling cones (SC10 container, Stuewe & Sons, Inc.) filled with sterile sand and perlite (1 : 1 volume) and inoculated with 10 mL of 10⁻¹ soil solution. The pots (4 cm × 20 cm) were placed for 8 wks in growth chambers at 20°C and 16 hrs light. At harvest, large red nodules were selected, surface sterilized for 1 min with 0.5% sodium hypochlorite, and a loop of inoculum was streaked on Yeast Extract Mannitol Agar (YEMA). Two nodules were collected from two different plants per soil, to produce a total of eight wild strains isolated from single colonies (Leb-M1, Leb-M2, Syr-M1, Syr-M2, Jord-M1, Jord-M2, Turk-M1, Turk-M2). Commercial strains of mesorhizobia were also provided from Novozyme Biologicals (NB-M4, NB-M5, NB-M6, NB-M7, NB-M8), Becker Underwood (BU-M3), and EMD (EMD-M1). Three varieties of US chickpeas (Myles, Troy, and Sierra) were supplied by the USDA-ARS Grain Legume Genetics-Physiology Research Unit, Pullman, WA, and three ME varieties (Arman, Almaz, and Hashem) were supplied from the U.S. National Plant Germplasm System, Regional Plant Introduction Station in Pullman, WA. Five days before seed inoculation, strains were grown in 1 X yeast extract broth at 22°C on an orbital shaker. Cells were collected by centrifugation at 4000 rpm for 4 min. An optical density () corresponding to a bacterial count of 10⁶–10⁷ was determined using a spectrophotometer at 640 nm wavelength. Inocula were diluted to 10⁵–10⁶ cfu per mL. One mL of wild (Leb-M1, Syr-M1, Jord-M1, Turk-M1) or commercial diluted inocula (NB-M4, NB-M5, BU-M3, EMD-M1) was added to each cone, which contained one chickpea seed. Arman was also inoculated with an additional four wild (Leb-M2, Syr-M2, Jord-M2, Turk-M2) and four commercial strains (NB-M6, NB-M7, NB-M8, EMD-M6).

Treatments were a factorial design of two main factors, rhizobial strain and crop variety. Varieties by strain pairings were grown in 164 mL seedling cones filled with sand-perlite mixture (1 : 1 volume) using five replicates. For precise measurement of biological N fixation, an isotopic 15-N method was used. Two conditions must exist for the effective use of the isotope dilution method for determining BNF: (1) the addition of N should be low and (2) the N addition should be thoroughly mixed in the medium [14–16].

Plants were irrigated with 10 mL 2 mM (¹⁵NH₄)₂ SO₄ (5% atom excess) three times a week for the first two weeks, and with H₂O thereafter [15]. Sterile N-free Hoagland solution was added once a week to provide all necessary nutrients except N. After 8 weeks, the whole plants were harvested and dried at 65°C, after which above-ground biomass (AGB) and below-ground biomass (BGB) were separated at the root crown, and weighed. Atom percent ¹⁵N/¹⁴N (A%) of combined AGB and BGB was measured by an isotope ratio mass spectrometer (Thermo-Finigan, Delta Plus Advantage). The atom % excess (A% − 0.366 A%) was used to calculate the proportion of N fixed (PNF): PNF = 1 − [% atom excess ¹⁵N in fixing crop/% atom excess ¹⁵N in nonfixing crop] × 100 [18]. Wheat was used as a control in the experiment as described by Knowles and Blackburn [19] and Smith et al. [20].

Five replicates were distributed in a randomized complete block design blocked by time. Analysis of variance (ANOVA) and correlations between various parameters were determined using SAS system for Windows version 9.1. Means were calculated with LSM in Proc Mixed. Data were considered significantly different at according to Tukey’s method in Proc Mixed.

2.2. PCR, Sequencing and Sequence Analysis

Genomic DNA was extracted with a Fast DNA SPIN KIT (MP Biomedicals Inc, OH) from seven commercial (EMD-M1, BU-M3, NB-M4, NB-M5, NB-M6, NB-M7, and NB-M8) and four wild (Leb-M1, Turk-M2, Syr-M2, and Jord-M1) isolates. In addition, DNA was extracted from two commercial strains of Rhizobium leguminoserum bv. viciae (NB-R2, NB-R7). Using the primer pair fD1 (5′-AGAGTTTGATCCTGGCTCAG-3′) and rD1 (5′-AAGGAGGTGATCCAGCC-3′), an approximately 1340 bp product specific to nearly full length of 16S rRNA gene was amplified by PCR [21]. Cycling conditions were as follows: one cycle of 95°C for 2 min followed by 30 cycles of 94°C for 40 s, 52°C for 40 s, and 72°C for 1 min. This was followed by a final extension at 72°C for 7 min. Amplicons were resolved on a 1.5% agarose (1 X TBE, 90 mM Tris pH 8.0, 90 mM boric acid, 2 mM EDTA) gel, stained with ethidium bromide, and visualized with UV light. The amplicons were purified using a QIAquick PCR Purification kit (QIAGEN Inc, CA) following manufacturer’s instructions, and the purified DNA samples were sequenced in both orientations using the ABI PRISM 377 DNA Sequencer (Applied Biosystems Inc, CA). Sequences were edited, and a consensus sequence for each amplicon was derived using Vector NTI Advance 11 (Invitrogen Inc, CA). Consensus sequences were compared with GenBank accessions by a BLAST search [22]. Multiple sequence alignments were performed by ClustalW [23]. Phylogenetic analyses by the neighbor-joining method and a consensus tree were generated using Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0 [24].

3. Results

3.1. Fixed N

Varieties and strains both had a major influence on the PNF. Averaged across all strains, Sierra had greater PNF than Arman, Hashem, and Myles (Table 1). Myles had a lower PNF than all other varieties. The selection of Sierra could increase PNF in the plant 13.2% over Myles. Varieties from the US and ME were statistically similar in PNF.

Averaged across all varieties, strain Jord-M1 had greater PNF than Syr-M1 (Table 2), but the difference was small. The Jord-M1 strain provided an average 84.7% PNF compared to 81.4% using Syr-M1. Strains from the US and ME provided statistically similar levels of PNF.

3.2. Above- and Below-Ground Biomass

Varieties substantially influenced above and below-ground biomass () (Table 1). Sierra and Troy had higher AGB than Arman, Hashem, and Myles. The AGB in Almaz was statistically similar to Sierra, Troy, and Arman and greater than Hashem and Myles (Table 1). This could be due to the larger seed size of these varieties. The average weight per seed in Sierra, Troy, and Almaz was, respectively 0.57 g, 0.60 g, and 0.50 g while Myles weighed 0.16 g (Table 1). Almaz, Arman, and Troy had greater BGB than Myles, Hashem, and Sierra. The BGB in Troy was statistically similar to Sierra, Almaz, and Arman but greater than Hashem, and Myles (Table 1). The ME varieties had significantly greater BGB than the US varieties, which could be directly or indirectly related to the breeding traits selected in the US and ME. A greater BGB may provide the ME varieties a greater or wider tolerance to adverse environmental factors such as water stress. The PNF was highly correlated to AGB (; ) and BGB (; ), indicating that generally the larger plants had a lower proportion of plant N supplied by the fertilizer.

Arman tests with four additional ME and four additional commercial strains did not reveal a significant difference among these strains on the measured parameters (data not shown). All commercial and ME strains used in this trial were infective and effective on Arman.

3.3. Sequence Analysis

Wild ME nodulators and commercial inoculant strains separated into distinct phylogenetic clades, which were also distinct from the R. leguminoserum outgroup (Figure 1). One commercial strain, NB-M5, was also distinct from all other isolates. A BLAST search of 16S rDNA sequences identified all commercial strains as M. ciceri and all wild ME symbionts as Burkholderia sp. Syr-M2 had 99% sequence identity with Burkholderia fungorum, Jord-M1 had 99% sequence identity with Burkholderia cepacia, and Turk-M2 and Leb-M1 were identified only as Burkholderia sp. Wild ME symbiont sequences were 97-98% similar among them, but only 79-80% similar to commercial strains of M. ciceri and 80-81% similar to R. leguminoserum (data not shown). This is the first report of Burkholderia that nodulates chickpea.

Figure 1

Phylogenetic analysis of 16S ribosomal RNA sequences of commercial and wild chickpea symbionts. Tree constructed using Neighbor-Joining implemented in MEGA4 program [17]. Bootstraps values (1,000 replicates) are given at the branch nodes; branches reproduced in less than 70% of bootstrap replicates are collapsed. Isolates NB-R2 and NB-R7 are an outgroup of commercial strains of Rhizobium leguminosarum bv. viciae. Isolates EMD-M1, BU-M3, NB-M4, NB-M5, NB-M6, NB-M7, and NB-M8 are commercial chickpea inoculant strains of M. ciceri. Isolates Leb-M1, Turk-M2, Syr-M2, and Jord-M1 are wild chickpea nodulating isolates from Middle Eastern soils.

4. Discussion

In this study we used a nonlegume crop as a control and isotope dilution methods to track the proportion of N fixed [19] via Fried and Middleboe’s equation [18]. The total N considered “fixed” in Fried and Middleboe’s equation actually originates from all nonfertilizer sources. There was a positive correlation () between the weight of the seeds and PNF in the harvested plant. An increase in the seed weight corresponded to an increase in PNF and suggests that ¹⁴N was supplied from the seeds. When seed N was subtracted from PNF as defined in Fried and Middleboe equation, only two varieties demonstrated true N fixation. Myles had 8.8% true PNF and Hashem had 14.5% true PNF. In this case, Fried and Middleboe’s equation for the determination of PNF that the plants derived from the atmosphere needs to be adjusted by subtracting the amount of N in the plant derived from the seeds.

The ability of Burkholderia sp. to symbiotically fix N on other legumes has been recognized recently. Bontemps et al. [25] tested 143 symbionts from 47 Brazilian wild species of Mimosa and found that 98% were Burkholderia sp. Two strains of Burkholderia caribensis had been isolated previously from Mimosa nodules [26]. Burkholderia tuberum isolated from a South African legume has been proved to contain nod and nif genes similar to Rhizobiaceae and showed a genetic response with bean and cowpea [27]. Burkholderia cepacia, which was isolated from Acacia decurrens and Ormosia nitida in Brazil, contained nod C and nif H genes [28]. Several studies have demonstrated Burkholderia sp. isolated from nodules of papilionoid and other mimosoid legumes are able to form nodules on selective variety of legumes [29, 30]. Burkholderia belongs to the betaproteobacteria class and carries similar nod genes to rhizobia, which are alphaproteobacteria. The transfer of nod genes to the new N-fixing bacteria may have occurred via lateral gene transfer [29, 31].

Other research has demonstrated a strong influence of plant variety on N fixation. Bello et al. [32] found 70% and 25% variability in kg N fixed ha⁻¹ among three soybean varieties at two sites. The survival and number of inocululated rhizobia are very critical for successful nodulation and yet limited efforts have been made to improve rhizobial inoculants [11]. Inoculated rhizobia strains may not compete well against resident rhizobia. Inoculation by commercial strains filled 5–20% of nodules on soybeans in studies by Caldwell and Vest [33] and Kuykendall and Weber [34]. In these trials, variety had a stronger influence on PNF than did strain, and all strains tested were effective on all varieties. Consequently, the more efficient strategy for BNF improvement may be in breeding crops that support greater N fixation rather than rhizobia strain selection.

5. Conclusion

Increased BNF by legume-bacterial associations can be used to further improve global crop production or reduce the need for synthetic N fertilizers. As the cost of fertilizers increases, the value of improving BNF also increases. Variety of chickpeas had a greater effect on variation in PNF than did the symbionts strain. This study indicates the possibility to improve the efficiency of N fixation through selection of chickpea varieties that host effective symbionts and have a high demand for N. Four newly isolated N-fixing chickpea symbionts from Middle Eastern soils were identified as Burkholderia sp. Studies are needed to investigate additional N-fixing bacteria beyond those found within the Rhizobiaceae. A new strain from Jordan, Jord-M1 contributed to the highest PNF. These findings suggest that variety selection may potentially improve PNF with a wide variety of symbionts, leading to higher yield, higher protein content, and/or more residual N for following crops.

Abbreviations

PNF:	Proportion of plant nitrogen provided by fixation
BNF:	Biological nitrogen fixation
N:	Nitrogen
ME:	Middle East
US:	United States
AGB:	Above-ground biomass
BGB:	Below-ground biomass.

References

FAO, Current World Fertilizer and Trends and Outlook to 2011-2012, FAO, Rome, Italy, 2008.
USDA-ERS, Average U.S. Farm Prices of Selected Fertilizers, 1960–2011, 2011.
IFA (International Fertilizer Industry Association), Greenhouse Gas Emissions and Fertilizer Production, 2009.
EIA (U.S. Energy Information Administration), World Proved Reserves of Oil and Natural Gas, Most Recent Estimates, 2009.
K. Albala, Beans: A History, Berg Publishers, New York, NY, USA, 2007.
M. R. Naghavi and M. R. Jahansouz, “Variation in the agronomic and morphological traits of Iranian chickpea accessions,” Journal of Integrative Plant Biology, vol. 47, no. 3, pp. 375–379, 2005.
View at: Publisher Site | Google Scholar
NASS National Agriculture Statistics Service, Quick Stats for Chickpeas (Garbanzo), 2011.
M. J. Unkovich and J. S. Pate, “An appraisal of recent field measurements of symbiotic N₂ fixation by annual legumes,” Field Crops Research, vol. 65, no. 2-3, pp. 211–228, 2000.
View at: Publisher Site | Google Scholar
C. Carranca, A. De Varennes, and D. Rolston, “Biological nitrogen fixation by fababean, pea and chickpea, under field conditions, estimated by the ¹⁵N isotope dilution technique,” European Journal of Agronomy, vol. 10, no. 1, pp. 49–56, 1999.
View at: Publisher Site | Google Scholar
J. Brockwell, R. R. Gault, D. F. Herridge, L. J. Morthorpe, and R. J. Roughly, “Studies on alternativemeans of legume inoculation: microbiological and agronomic appraisals of commercial procedures for inoculating soybeans with Bradyrhizobium japonicum,” Australian Journal of Agricultural Research, vol. 40, pp. 753–762, 1989.
View at: Google Scholar
J. Brockwell and P. J. Bottomley, “Recent advances in inoculant technology and prospects for the future,” Soil Biology and Biochemistry, vol. 27, no. 4-5, pp. 683–697, 1995.
View at: Google Scholar
R. Abi-Ghanem, L. Carpenter-Boggs, and J. L. Smith, “Cultivar effects on nitrogen fixation in peas and lentils,” Biology and Fertility of Soils, vol. 47, pp. 115–120, 2011.
View at: Publisher Site | Google Scholar
C. P. Horn, R. C. Dalal, C. J. Birch, and J. A. Doughton, “Nitrogen fixation in chickpeas as affected by planting time and tillage practice,” in Proceedings of the 8th Australian Agronomy Conference, D. L. Michalk and J. E. Pratley, Eds., pp. 1–5, Queensland, Australia, February 1996.
View at: Google Scholar
P. M. Chalk, “Estimation of N₂ fixation by isotope dilution: an appraisal of techniques involving ¹⁵N enrichment and their application,” Soil Biology and Biochemistry, vol. 17, no. 4, pp. 389–410, 1985.
View at: Google Scholar
S. K. A. Danso, G. Hardarson, and F. Zapata, “Misconceptions and practical problems in the use of ¹⁵N soil enrichment techniques for estimating N₂ fixation,” Plant and Soil, vol. 152, no. 1, pp. 25–52, 1993.
View at: Publisher Site | Google Scholar
F. R. Warembourg, “Nitrogen fixation in soil and plant systems,” in Nitrogen Isotope Techniques, R. Knowles and T. H. Blackburn, Eds., pp. 127–155, Academic Press, New York, NY, USA, 1993.
View at: Google Scholar
K. Tamura, M. Nei, and S. Kumar, “Prospects for inferring very large phylogenies by using the neighbor-joining method,” Proceedings of the National Academy of Sciences of the United States of America, vol. 101, no. 30, pp. 11030–11035, 2004.
View at: Publisher Site | Google Scholar
M. Fried and V. Middelboe, “Measurement of amount of nitrogen fixed by a legume crop,” Plant and Soil, vol. 47, no. 3, pp. 713–715, 1977.
View at: Publisher Site | Google Scholar
R. Knowles and H. T. Blackburn, Nitrogen Isotope Techniques, Academic Press, San Diego, Calif, USA, 1993.
S. C. Smith, D. F. Bezdicek, R. F. Turco, and H. H. Cheng, “Seasonal N₂ fixation by cool-season pulses based on several ¹⁵N methods,” Plant and Soil, vol. 97, no. 1, pp. 3–13, 1987.
View at: Publisher Site | Google Scholar
W. G. Weisburgh, S. M. Barns, D. A. Pelletier, and D. J. Lane, “16S ribosomal DNA amplification for phylogenic study,” Journal of Bacteriology, vol. 173, pp. 697–703, 1991.
View at: Google Scholar
S. F. Altschul, W. Gish, W. Miller, E. W. Myers, and D. J. Lipman, “Basic local alignment search tool,” Journal of Molecular Biology, vol. 215, no. 3, pp. 403–410, 1990.
View at: Publisher Site | Google Scholar
J. D. Thompson, D. G. Higgins, and T. J. Gibson, “CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice,” Nucleic Acids Research, vol. 22, no. 22, pp. 4673–4680, 1994.
View at: Google Scholar
K. Tamura, J. Dudley, M. Nei, and S. Kumar, “MEGA4: molecular evolutionary genetics analysis (MEGA) software version 4.0,” Molecular Biology and Evolution, vol. 24, no. 8, pp. 1596–1599, 2007.
View at: Publisher Site | Google Scholar
C. Bontemps, G. N. Elliott, M. F. Simon et al., “Burkholderia species are ancient symbionts of legumes,” Molecular Ecology, vol. 19, pp. 44–52, 2010.
View at: Google Scholar
K. Vermis, T. Coenye, J. J. Lipuma, E. Mahenthiralingam, H. J. Nelis, and P. Vandamme, “Proposal to accommodate Burkholderia cepacia genemovar VI as Burkholderia dolosa sp. nov,” International Journal of Systematic and Evolutionary Microbiology, vol. 54, pp. 689–691, 2004.
View at: Google Scholar
G. N. Elliott, W. M. Chen, C. Bontemps et al., “Nodulation of Cyclopia spp. (Leguminosae, Papilionoideae) by Burkholderia tuberum,” Annals of Botany, vol. 100, no. 7, pp. 1403–1411, 2007.
View at: Publisher Site | Google Scholar
P. Menna, M. Hungria, F. G. Barcellos, E. V. Bangel, P. N. Hess, and E. Martínez-Romero, “Molecular phylogeny based on the 16S rRNA gene of elite rhizobial strains used in Brazilian commercial inoculants,” Systematic and Applied Microbiology, vol. 29, no. 4, pp. 315–332, 2006.
View at: Publisher Site | Google Scholar
L. Moulin, A. Munive, B. Dreyfus, and C. Boivin-Masson, “Nodulation of legumes by members of the β-subclass of Proteobacteria,” Nature, vol. 411, no. 6840, pp. 948–950, 2001.
View at: Publisher Site | Google Scholar
C. F. Barrett and M. A. Parker, “Prevalence of Burkholderia sp. nodule symbionts on four mimosoid legumes from Barro Colorado Island, Panama,” Systematic and Applied Microbiology, vol. 28, no. 1, pp. 57–65, 2005.
View at: Publisher Site | Google Scholar
A. Sy, E. Giraud, P. Jourand et al., “Methylotrophic Methylobacterium bacteria nodulate and fix nitrogen in symbiosis with legumes,” Journal of Bacteriology, vol. 183, no. 1, pp. 214–220, 2001.
View at: Publisher Site | Google Scholar
A. B. Bello, W. A. Ceron-Dias, C. D. Nickell, E. O. Elsheriff, and L. C. Davis, “Influence of cultivar, between-row spacing and plant population of fixation of soybeans,” Crop Science, vol. 20, pp. 751–775, 1980.
View at: Google Scholar
B. E. Caldwell and G. Vest, “Effects of Rhizobium japonicum strains on soybean yields,” Crop Science, vol. 10, pp. 19–21, 1970.
View at: Google Scholar
L. D. Kuykendall and D. F. Weber, “Genetically marked Rhizobium identifiable as inoculum strain in nodules of soybean plants grown in fields populated with Rhizobium japonicum,” Applied and Environmental Microbiology, vol. 36, pp. 15–919, 1978.
View at: Google Scholar

Copyright

Copyright © 2012 Rita Abi-Ghanem et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

PDF Download Citation

Download other formats

Order printed copies

Views

3660

Downloads

1560

Citations