Isolation and selection of nitrogen-fixing bacteria

In October 2014, 10 healthy and vigorous Bermuda grass plants growing ruderally were selected and extracted in the Department of Agriculture and Livestock of the University of Sonora located at 29° 00’ 51.72” North latitude and 111° 08’ 01.10” West longitude, with an altitude of 149 m, in this site the climate is classified as BW (h’) hw (e’), which corresponds to the category of very arid, extreme and warm. The texture of the soil at the collection site is sandy loam. The aerial part of each plant was eliminated and approximately 1 kg of soil containing the roots was collected.

The soil was removed and the roots were washed with running water. The roots were cut into approximately 1 cm segments and rinsed 3 times for 10 min with sterile distilled water. Three root pieces were placed in screw-capped glass tubes, which contained 15 mL of semi-solid NFb medium (^{Döbereiner and Day, 1976}) nitrogen-free. The tubes were closed, sealed with parafilm and incubated at 28 °C for 5 days. 1 mL of the microbial mass developed on the surface of the medium was collected and serial dilutions were seeded in Petri dishes containing solid NFb medium (agar, 15 g L^-1).

After one week of incubation, the change from blue to green color around the colony was indicative that the isolate had nitrogen-fixing activity (^{Baldani et al., 2014}). Twelve isolates with different morphologies were obtained and two of them, 0710 and 0711, were chosen to continue with phenotypic and biochemical characterization and molecular identification. The selection criteria were: greater intensity and speed in the change of color. The selected isolates were planted in Luria-Bertani medium (LB) for their conservation and subsequent analysis.

Phenotypic and biochemical characterization

Isolates 0710 and 0711 were cultured in total count agar for morphological analysis and characterized by the biochemical tests and carbon assimilation profiles of the API 20 E System commercial kit (BioMerieux Laboratory, France).

Molecular identification by sequencing a 16S rRNA gene fragment

The total DNA extraction of the selected isolates was carried out with the phenol-chloroform technique (^{Ausubel et al., 1994}). The DNA was amplified by means of the polymerase chain reaction (PCR) using the F2C universal primers (5’-AGA GTT TGA TCA TGG CTC-3’) and C (5-ACG GGC GGT GTG TAC-3’), which amplify a fragment of around 1 600 bp. The reaction mixture contained: 20 ng of total DNA, 12.5 μL of GoTaq^® Green Master Mix enzyme (Promega), 0.5 μM final concentration of each primer and water-free nucleases until a final volume of 25 μL was completed. The PCR conditions were: 1) initial denaturation at 95 °C for 2 min; 2) amplification, consisting of 30 cycles with the following steps: a) denaturation at 95 °C for 1 min; b) alignment at 55 °C for 1 min; and c) extension at 72 °C for 2 min and finally; and 3) final extension at 72 °C for 5 min. The BioRad C1000 thermocycler was used.

A second amplification was performed using the primers U1: 5’-CCA GCA GCC GCG GTA ATA CG-3’ and C, in order to obtain a smaller fragment (~ 900 bp), suitable for sequencing. As template DNA, 1 μL of the PCR products of the first amplification reaction was used. The amplification program was the same as the previous one. The final products were purified with a Wizard^® SV Gel and PCR Clean-Up System kit (Promega Co.) and sequenced in both directions using again the U1 and C primers separately, with an ABI PRISM^® 3730 Genetic Analyzer (Applied Biosystems). The sequences were compared with those of the NCBI GenBank database using the BLAST algorithm and aligned using the ClustalOmega program.

In vitro test of biological nitrogen fixation (FBN)

Isolates 0710 and 0711 were cultured in liquid enriched medium (1.5 g of meat peptone, 3 g of yeast extract, 6 g of gelatin peptone and 1 g of glucose in 1 000 mL of distilled-deionized water). A culture sample was taken and adjusted to a concentration between 10⁵ and 10⁶ CFU mL^-1. 100 μL of the adjusted sample of each isolate was taken and inoculated in triplicate into 125 mL flasks with 50 mL of N-free thiamine-biotin medium. The flasks were incubated at 30 °C under agitation for 7 days.

The capacity of FBN was estimated according to ^{Kanimozhi and Panneerselvam (2010)}, by determining the ammonia produced during the digestion process. 2 mL of the culture was placed in a Microkjeldahl flask, 2 mL of H₂SO₄ (d= 1.84), 1 mL of H₂O₂ (30%) and 1 g of digester mixture (100 g of Na₂SO₄ + 10 g of CuSO₄5H₂O + 1 g of selenium) were added, at 350-375 °C and digested by boiling for 1.5 h. The digestion products were separated by distillation with 10 mL of NaOH 10 M and the ammonia produced was determined by titration with 0.0025 HCl M.

Qualitative phosphate solubilization assay

The phosphate solubilization test was carried out according to ^{Malboobi et al. (2009)}. A needle tip was immersed in medium enriched with isolates 0710 and 0711 and was inoculated by stinging in the center of Petri dishes with the solid medium of Pikovskaya (15 g of agar, 10 g of glucose, 5 g of Ca₃ (PO₄)₂, 0.5 g of yeast extract, 0.5 g of (NH₄)₂SO₄, 0.2 g of KCl, 0.1 g of MgSO₄7H₂O, 0.1 mg of MnSO₄2H₂O, 0.1 mg of FeSO₄ in 1000 mL of distilled water). Five boxes were planted per isolate and incubated at 30 °C for 7 days. The development of a clear zone around the colony was the sign of solubilization of inorganic phosphate.

Solubilization of inorganic phosphate (IPS)

Aliquots of 1 mL of cultures 0710 and 0711 were taken in liquid enriched medium and placed in 3 Erlenmeyer 500 mL flasks for each bacterium containing 200 mL of sterile Pikovskaya liquid medium. The flasks were kept under agitation (300 rpm) at 30 °C for 2 days. Sterile medium was used without inoculation as a negative control. After incubation, 10 mL of each flask was taken for centrifugation. The determination was carried out as described by ^{Pote and Daniel (2000)}.

5 mL of each supernatant was taken and mixed with 800 μL of mixed reagent (50 mL of 2.5 M H₂SO₄, 15 mL of 4% ammonium molybdate, 30 mL of 0.1 M ascorbic acid and 5 mL of potassium tartrate solution 0.004 M) to quantify the release of phosphorus to the medium. A standard curve was prepared with known concentrations (0.15, 0.3, 0.6, 0.9, 1.2 and 1.5 μg mL^-1), prepared from a stock solution containing 50 μg mL^-1 of P. The absorbance of standard solutions was read and samples at 880 nm in a Spectronic 21D manual spectrophotometer (Milton Roy), using a reagent blank as a reference solution. The samples were proportionally diluted for reading in the graphic range.

Assay of the enzymatic activity of acid phosphatase (ACF) and alkaline (ALF)

The enzymatic activity of acid phosphatase and alkaline phosphatase was determined in triplicate using the modified test of ^{Juma and Tabatabai (1988)}. A mixture of 1 mL of the supernatant obtained in the inorganic phosphate solubilization test of isolates 0710 and 0711 was made with 1 mL of 25 mM p-nitrophenyl phosphate (pNPP) and 4 mL of modified universal buffer [12.1 g of tris (hydroxymethyl) aminomethane, 11.6 g of maleic acid, 14 g citric acid and 6.3 g of H₃BO₃ in 488 ml of 1 M NaOH by 1 liter with distilled-deionized water. This buffer was adjusted to the desired pH and stored at 4 °C. Before use, it was diluted five times with distilled-deionized water]. The pH values of the buffer were 6.5 and 11, for the acid or alkaline phosphatase assay, respectively.

Each reaction mixture was incubated at 37 °C and after 1 h of incubation, 1 mL of CaCl₂ 0.5 M and 4 mL of NaOH 0.5 M were added to finish the reaction. The assay mixtures were filtered and their absorbance at 410 nm was recorded in a Spectronic 21D manual spectrophotometer (Milton Roy) to quantify the intensity of the yellow color due to the formation of p-nitrophenol (pNP). Quantification was performed using a pNP standard of known concentration and the results were expressed in units of activity (U), defining 1 U as the amount of enzyme required to release 1 μg of pNP in 1 mL of culture filtrate at the conditions of the assay (^{Prasanna et al., 2011}).

ACC-deaminase assay

The activity of the ACC-deaminase enzyme was determined in triplicate according to the method developed by ^{Penrose and Glick (2003)}. This assay determines the amount of (-ketobutyrate produced when the enzyme breaks down the 1-aminocyclopropane-1-carboxylic acid molecule (ACC). The number of μmoles of α-ketobutyrate produced in the reaction is determined by comparing the absorbance of a sample at 540 nm in a Spectronic 21D manual spectrophotometer (Milton Roy), against a standard α-ketobutyrate curve prepared in a range of 0.1 and 1 μmol.

Production of Indolaacetic acid (AIA)

It was carried out according to the method of ^{Loper and Scroth (1986)} and was carried out both in the absence and in the presence of 500 μg mL^-1 of tryptophan. Previously bacterial isolates 0710 and 0711 were cultured in 3 mL of medium enriched for 24 h at 28 °C and 200 rpm. 20(L of that culture were transferred to 3 tubes with 5 mL of new enriched media for each isolate, without supplementation or supplemented with 500 μg mL^-1 of tryptophan, incubated for 72 h at 28 °C and 200 rpm. After this time, the cultures were centrifuged for 10 minutes at 10 000 xg, 1 mL of supernatant was taken and mixed with 50 μL of orthophosphoric acid and 2 mL of the Salkowsky reagent (1 mL of 0.5 M FeCl₃ in 50 mL of 35% HCIO₄), incubating at 28 °C for 30 min.

At the end of that period, the appearance of a pink color indicated the presence of AIA. The quantification of AIA was performed by measuring the absorbance at 530 nm in a Spectronic 21D manual spectrophotometer (Milton Roy) with the help of a standard graph of AIA (Sigma) obtained in the range of 0 to 40 μg mL^-1 (^{Ahmad et al., 2008}).

Production of gibberellic acid (AG₃)

Isolates 0710 and 0711 were cultured separately in tubes with 3 mL of enriched liquid medium, and incubated overnight at 28 °C and 200 rpm. 20 μL of each culture were transferred to 3 flasks of 50 mL of medium enriched per isolate and incubated under the same conditions for 72 h. The samples were centrifuged to remove the cell biomass. The determination was made according to ^{Berrios et al. (2004)}. 1 mL of the supernatant was placed in a 10 mL volumetric flask and acidified by gauging with 3.75 M HCl. It was stirred vigorously for 10 s and allowed to react for 45 min to convert the gibberellic acid into giberelenic acid.

This compound was extracted once with 30 mL of ethyl acetate. The aqueous phase was discarded and the ethyl acetate phase was measured by a Milton Roy Spectronic 21D spectrophotometer (Milton Roy) at 254 nm. To estimate the amount of gibberellic acid produced, a calibration chart was prepared from standard solutions of: 50, 75, 100, 120, 150, 200, 240, 300, 400, 500 and 600 μg mL^-1, prepared at from a stock solution of 100 mg AG₃ in 250 mL of absolute ethanol. The solutions were acidified and extracted with ethyl acetate, similar to the sample. The values of absorbance against concentration were plotted and adjusted by linear regression.

Production of siderophores

For this test, cultures 0710 and 0711 were diluted in peptone water and planted at three equidistant points of three Petri dishes with agar-chromium azurol S (CAS) for each culture. The boxes were incubated at 28 °C for 48 h. After that time the siderophore producing bacteria, capable of fixing the iron, will produce an orange halo around the colonies in the blue medium (^{Schwyn and Neilands, 1988}).

Production of hydrocyanic acid (HCN)

Bacteria 0710 and 0711 were cultured on glycine-modified King B agar medium (4.4 g L^-1). A 8 cm diameter disc of Whatman filter no. 1 paper was placed inside the lid of the Petri box, previously soaked with a sterile solution of 1% picric acid in 10% sodium carbonate (filter paper and picric acid were sterilized separately). The plates were sealed with parafilm and incubated at 28 °C for 48 h. Three boxes were seeded by bacteria. The development of a reddish-brown color on the filter paper would indicate that the samples are positive for the production of HCN (^{Ahmad et al., 2008}).

Antifungal activity assay

Isolates 0710 and 0711 were tested to determine the inhibitory effect of growth on the mycelial growth of Ceratobasidium sp., Fusarium oxysporum, F. falciforme and F. brachygibbosum, all of them previously isolated as causes of root rot in watermelon plants in Sonora, Mexico (^{Rentería-Martínez et al., 2018}). The test was carried out on the complex agar medium (10 g of peptone, 40 g of D-glucose, 10 g of yeast extract, 18 g of agar in 1000 ml of water). The fungi were grown on potato dextrose agar (PDA). The pure bacterial cultures were cultured in liquid medium enriched at 28 °C for 24 h. 100 μL of each bacterial culture was spread on the surface of the complex agar.

Then, an 8 mm PDA disc covered with mycelium of each fungus was placed in the center of the Petri dish. At the same time, control experiments were performed, placing a disk of each fungus in the middle of the complex agar without bacteria. Three boxes were planted by bacteria. The diameter of each fungal colony was measured after an incubation period of 7 days at 28 °C. The inhibitory effect was calculated as the percentage of the inhibition rate: IR%= (CB)(100)/C, where C is the diameter of the fungal mycelium control and B the diameter of the mycelium grown in the presence of bacteria (^{Laslo et al., 2012}).

Statistical analysis

The analysis of the results was carried out using the t test (for values of two independent variables), with a level of significance of 0.05%, using the IBM SPSS Statistics 24.0.0 software. Figure 1 referring to the inhibitory effect of bacterial isolates on pathogenic fungi was also performed analyzing the data with the same software.

Figure 1 Inhibitory activity of Pm0710 and Cm0711 against the growth of phytopathogenic fungi isolated from watermelon plants with symptoms of root rot.  

Antifungal capacity

Cm0711 showed significantly higher levels of inhibition than Pm0710 in all tested pathogens. C. massiliae had inhibition rates of around 60 to 80% in all fungi. P. monteilii had an effect mainly on Fusarium brachygibbosum and F. solani (55% and 40%, respectively) and less on Ceratobasidium sp. and F. oxysporum, 6 and 20% respectively (Figure 1).

The biological control of phytopathogens is the main indirect mechanism for promoting the growth of plants by rhizobacteria. In general, competition for nutrients, exclusion of niches, induced systemic resistance and production of antifungal metabolites are the main modes of biological control activity of PGPR. Antifungal metabolites include antibiotics, enzymes that degrade the cell wall, chelating siderophores of Fe and HCN (^{Sindhu and Dadarwal, 2001}; ^{Ahmad et al., 2008}; ^{Ravindra et al., 2008}; ^{Bhattacharyya and Jha, 2012}; ^{Glick, 2012}).

In our experiment, both Cm0711 and Pm0710 inhibited the growth of the four phytopathogens tested, however, the inhibitory effect of Cm0711 was significantly higher. The mechanisms by which these isolates exert control over plant pathogenic fungi remain unknown and should be elucidated in later work. Although it is known that some species of Chryseobacterium can act as PGPR (^{Shin et al., 2007}; ^{Dardanelli et al., 2010}; ^{Montero-Calasanz et al., 2013}), there are no reports on the potential characteristics as a promoter of C. massiliae.

With respect to Pseudomonas monteilii, it has been observed that certain isolates can associate with rice plants and promote their growth due to their ability to fix nitrogen. (^{Habibi et al., 2014}), while others significantly improve the length of the primary root, the outbreak and the number of lateral roots of the soybean sprouts (^{Wahyudi and Astuti, 2011}). The phosphate solubilization capacity of P. monteilii strains has been previously reported (^{Ravindra et al., 2008}; ^{Dharni et al., 2014}). ^{Pandya and Desai (2014)}, found thirteen isolates capable of synthesizing gibberellic acid, in other works P. monteilii caused the inhibition of the growth of Sclerotium rolfsii by the production of diffusible antibiotics, volatile metabolites, hydrogen cyanide, siderophores and proteases (^{Rakh et al., 2011}).

Another isolate produces 2,4-di-tert-butylphenol, an active volatile organic compound that acts as an inhibitor of β-tubulin in Fusarium oxysporum (^{Dharni et al., 2014}).