Biological Efficacy of Plant Growth–Promoting Bacteria and Arbuscular Mycorrhizae Fungi: Assessments in Laboratory and Greenhouse Conditions

Abstract

Utilizing the interactions of microorganisms with plants offers a favorable path to increase crop production and replace the use of synthetic fertilizers. Different bacteria and fungi have been used as biofertilizers to improve agricultural production, yield, and sustainability. Beneficial microorganisms can act as free‐living organisms, symbiotes, and endophytes. Soil bacteria called plant growth–promoting bacteria (PGPB) and fungi called arbuscular mycorrhizae fungi (AMF) stimulate the growth and health of plants by direct and indirect mechanisms including nitrogen fixation, phosphorus solubilization, phytohormone production, enzyme production, antibiotic synthesis, and induced systemic resistance. To use these microorganisms as a biofertilizer, it is necessary to assess their efficacy under laboratory and greenhouse conditions. Few reports detail the methods used to develop a test under different environmental conditions, and without these details it is difficult to develop suitable methodologies to evaluate microorganism‐plant relationships. We describe four protocols that go from sample preparation to testing in vitro the efficacy of different biofertilizers. Each protocol can be used to test a different biofertilizer microorganism, focusing on bacteria such as Rhizobium sp., Azotobacter sp., Azospirillum sp., Bacillus sp. as well as AMF such as Glomus sp. These protocols can be used in several stages of biofertilizer development, including microorganism selection, microorganism characterization, and in vitro evaluation of efficacy for the registration process. © 2023 Wiley Periodicals LLC.

Basic Protocol 1: Evaluating the biological effect of biofertilizer based on PGPB under laboratory conditions

Basic Protocol 2: Evaluating the biological effect of biofertilizer based on PGPB under greenhouse conditions

Basic Protocol 3: Evaluating the biological effect of biofertilizer based on symbiotic nitrogen‐fixing bacteria

Basic Protocol 4: Evaluating the biological effect of biofertilizer based on AMF

INTRODUCTION

For years, chemical fertilizers based on nitrogen, phosphorus, or potassium have been applied excessively to increase agricultural productivity (Kumar et al., 2022). Only 30%‐40% of these nutrients are absorbed by plants due to low fertilizer‐use efficiency, causing environmental pollution including heavy metal accumulation in soil and eutrophication of water sources (Basu et al., 2021). In order to resolve these problems, sustainable and eco‐friendly technologies are needed that could reduce the use of synthetic fertilizers. A biofertilizer is defined as a product based on culturable microorganisms, including bacteria and fungi, that are isolated from soil or plant rhizospheres and can promote plant growth when applied to seeds, plants, or soil (Sharma et al., 2021; Tiwari et al., 2022). Using such bioproducts can ensure chemical‐free and environmentally friendly farming practices in organic farming and sustainable agriculture.

Biofertilizers can be classified according to their functions and modes of action (Mahanty et al., 2017). These microorganisms can be free‐living in the soil or have symbiotic and other endophytic associations with plants (Thomas & Singh, 2019). Plant growth–promoting bacteria (PGPB) and arbuscular mycorrhizae fungi (AMF) stimulate plant growth and health by both direct and indirect mechanisms. Direct mechanisms refer to traits that result in direct promotion of plant growth. This could be by providing beneficial compounds to host plants, facilitating nutrient uptake from the soil (e.g., N, P, K, Fe, Zn, and S), and modulating phytohormones (e.g., abscisic acid, cytokinin, ethylene, indole acetic acid) and gibberellin, a relevant factor for endophytes that are candidates for biostimulant products in agricultural crops. Indirect mechanisms refer to traits that inhibit the function of one or more plant pathogenic organisms (Orozco‐Mosqueda et al., 2021). Examples include bacterial siderophores, bacterial enzymes (e.g., chitinases, cellulases, and β‐1,3‐glucanases), and induced systematic resistance (Rath et al., 2021). The genera most used for biofertilizer production are bacteria including Rhizobium sp., Azotobacter sp., Azospirillum sp., Pseudomonas sp., Bacillus sp., and AMF including Glomus sp., Acaulospora sp., Gigaspora sp., Sclerocystis sp., Entrophospora, and Scutellospora sp. (Nosheen et al., 2021).

For biofertilizer to be developed into a high‐quality product, extensive research and proper experimental design are required. Five fundamental procedures are involved in development of a biofertilizer. As a first step, microorganisms associated with plants need to be isolated and identified. The selected strain must be competitive and highly efficient toward indigenous populations present in the soil; if it is preferable to use a combination of strains, both must be compatible with developing an efficient inoculum (Sharma et al., 2021; Raimi et al., 2021). Functional characterization of candidate strains is then performed using common laboratory techniques, including differential media or qualitative testing. The next step involves cultivating microorganisms to prepare starter cultures and inocula, as well as preparing carrier materials and incorporating the microorganisms into them (Vassilev et al., 2015). The viability of the reference strain while maintaining the inherent characteristics of the strain of interest is important for effective performance under greenhouse or field conditions. Optimal conditions are achieved by monitoring microbial growth curves under different conditions. Typically, propagation of strains is achieved using conventional fermenter systems. The two main types of fermentation used to produce biofertilizers are solid‐state and submerged fermentation (Suthar et al., 2017). Prototyping is the next step and involves testing different product shapes. Four dispersal forms are generally used: dry inoculant (mostly powders), slurry (a suspension of powder in liquid), granular, and liquid (Sharma et al., 2021; Rath et al., 2021). Carrier materials can be organic, inorganic, or synthetic with appropriate physicochemical properties for maintaining the microbial strains in good physiological condition under storage and on the field. Finally, quality control (QC), packaging, and storing biofertilizers based on carrier matrices are performed (Sarbani & Yahaya, 2022).

The recovery and multiplication techniques for AMF differ from those for other microorganisms such as bacteria and yeast, especially due to the symbiotic relationship with a host plant. AMF are multiplied by monoxenic culture, a regulated method of multiplication in which AMF are grown in a host plant (Basiru et al., 2021; Raimi et al., 2021). Multiplication is done in a pothouse under controlled conditions for commercial production (greenhouse). This technique enables accurate monitoring of inoculum spores and mycelia associated with host plant roots. Critical factors in the generation of AMF inocula include soil nutrient conditions and a high degree of host and AMF synergy (Salomon et al., 2022). The host plant must be able to grow quickly with significant root development and should have a favorable AMF colonization affinity. Monoxenic culture offers the benefit of producing a contamination‐free biofertilizer propagule (Raimi et al., 2021; Salomon et al., 2022).

One of the challenges in biofertilizer production is quality control, which is extremely important and should be consistently attained in the industry. The criteria used to determine quality of a specific strain are limited to viability, conservation, purity, and bioassay under controlled conditions (biological efficacy). This last parameter must be validated using a standardized in vivo or in vitro bioassay that provides information about effects on the target plant and its interaction with the PGPB or AMF. In this context, in vitro means the test is performed in plants grown hydroponically or using a controlled substrate in pots, growth chambers, or laboratory conditions. In vivo means the test is performed in soil in either greenhouse or field conditions. The bioassay must have a low cost and a quantifiable response (effectiveness) (Salomon et al., 2022). It must also be repeatable and reliable in order to determine the effectiveness with accuracy. Variations in components may cause variability in the results and make the bioassay difficult to replicate. Currently, information on standardized bioassays is limited. Few reports describe the methodology of developing bioassays for specific purposes and, without these details, it is difficult to develop and apply bioassay methodologies for analyzing interactions between beneficial microorganisms and plants. Hence, there is an urgent need to adopt universal, highly reproducible methods to evaluate commercial biofertilizers that can be applied to various steps of development such as strain characterization, evaluation of prototypes, and final product quality.

Here, we present four protocols for assessing biological activity (effectiveness) of PGPB and AMF biofertilizers under laboratory and greenhouse conditions. These protocols describe and provide a workflow to assess the biological activity of biofertilizers before field application. Basic Protocols 1‐3 focus on PGPB (Fig. 1). The objective of each protocol depends on the microorganism of interest. Basic Protocol 4 presents a method for assessing mycorrhizal inoculants (Fig. 1). The proposed bioassay was designed to evaluate two mycorrhizal species (e.g., Glomus sp. and Claroideoglomus sp.) with a select host plant in sterilized substrates under controlled conditions. This approach is appropriate for efficient investigation of the biological activity of biofertilizers, with the outcome of developing a novel bioassay that is replicable in future work. With these methods, users and especially QC laboratories can obtain a variety of data parameters for decision‐making on the selection and characterization of a microorganism and in vitro evaluation of efficacy for the registration process. The data obtained are divided into two types: (1) agronomic response variables (e.g., weight, length, leaf area) that describe the response of units to the experimental treatments and (2) effectiveness expressed as a percentage.

Figure 1.

Protocols for assessing the biological efficacy of PGPB and AMF.

CAUTION: These protocols are designed to be used with biofertilizer microorganisms such as PGPB and AMF. All standard microbiological practices should be followed when working with these organisms. Information about the biosafety level of each specific microorganism should be well‐understood by the user. As examples used in these protocols, Azotobacter sp., Bacillus amyloliquefaciens, Rhizobium sp., and Glomus sp. are all Biosafety Level 1 (BSL‐1) organisms for which standard microbiological practices can be used. Biosafety tests such as pathogenicity, toxicity, and allergenicity assays have been widely performed for some Bacillus sp. strains and have shown their low risk to laboratory personnel and the environment.

NOTE: For all protocols, it is important to minimize the risk of sample contamination. All solutions and equipment coming into contact with biofertilizer and microbial strains must be sterile. All soil must be autoclaved prior to use (we recommend three cycles of sterilization at 121°C, 20 PSI, for 1 hr). Where indicated, steps must be performed in a biological safety cabinet (e.g., ESCO, SC2‐4E2) using appropriate sterile technique.

Basic Protocol 1. EVALUATING THE BIOLOGICAL EFFECT OF BIOFERTILIZER BASED ON PGPB UNDER LABORATORY CONDITIONS

Diazotrophic and non‐diazotrophic bacteria have different functionalities due to the different benefits to the development of different plant species by direct or indirect mechanisms (Glick, 2012). Among these benefits is the cycling of nutrients such as nitrogen and phosphorus in the soil, which helps increase their availability for their capture and use in plant nutrition (Adesemoye & Kloepper, 2009). Additionally, biofertilizers can improve plant growth through mechanisms involving the production and synthesis of phytohormones such as auxins, abscisic acid, cytokinins, ethylene, and gibberellins. Other benefits include the solubilization of phosphates, production of siderophores, and intrinsic systemic resistance to biotic stress, drought, sodium salinity, and heavy metals (Malusá & Vassilev, 2014). Some microbial species such as Azospirillum brasilense and Azotobacter chroococcum, which are free‐living in the soil, colonize the rhizosphere or plant tissue to positively affect plant growth and are used in biological inoculants (Domingues Duarte et al., 2020; Malusá y Vassilev, 2014).

Biological activity in a laboratory or greenhouse is one of the most important quality parameters in the production of microbial inoculants because it defines the efficacy of the bioproduct and the stability of the biological activity of the active ingredient (Hermann et al., 2015). In general, this parameter can be evaluated using field trials to determine production yield, plant growth, and nutrient absorption (Ohyama & Pham, 2006). The evaluation is carried out in legumes because symbiotic bacterial nitrogen fixers and non‐symbiotic bacteria are known to promote plant growth in legumes (Casas et al., 2019).

This protocol outlines how to quantify the effectiveness of response variables and can be evaluated in 15 days, which allows for a reduction in time to results. The effectiveness of biological inoculants based on growth‐promoting bacteria is assessed to determine the capacity of the bioproduct to generate differences in root length, aerial length, aerial dry weight, root dry weight, and foliar area. The test is conducted using negative control or untreated plants, where temperature, humidity, and photoperiod are controlled and external factors can be eliminated. At least three treatments are evaluated: a biofertilizer to be tested, a positive control (reference strain), and a negative control (sterile distilled water). Each treatment has ten experimental units and three replicates (see Statistical Analysis).

Materials

• 70% (v/v) ethanol (for sterilizing surfaces and supplies)

• Azotobacter chroococcum (ATCC 9043) or Azospirillum brasilense (ATCC 29145)

• Ashby's mannitol broth or DYGS broth (see recipes)

• Distilled water

• Soybean seeds (Glycine max; e.g., variety Taluma, Primavera, or Achagua)

• Biofertilizer: any PGPB‐based prototype or final product

• Autoclaved soil (Andisol, pH 4.0‐6.0, P 0.86 ppm, MO >10%, N 0.53%, K 0.1‐0.4 cmol+ kg^–1, Al 0.60 cmol+ kg^–1, Mg <1.5 cmol+ kg^–1, S <10 mg kg^–1, Ca (<3 cmol+ kg^–1) (Callejas et al., 2018; García Guzmán et al., 2019; Medina et al., 2019)

• Hoagland's nutrient solution (see recipe) without nitrogen

• Sterile 100‐ and 500‐ml beakers (e.g., Brand, BR90636)

• Parafilm

• 28°C shaking incubator (e.g., Labcompanion, SI‐300)

• Spectrophotometer (e.g., Beckman, DU530)

• Sterile forceps (e.g., Brand, T5790)

• Sterile absorbent paper (Wypall 80, SAP 30163165)

• 9‐oz plastic cups (Darnel, D630502A)

• Wypall disposable wipes

• Flexometer (e.g., Stanley, 52767)

• Leaf area meter (e.g., Marconi, AM350) (optional)

• Paper bags (e.g., Kraft, 20 × 9 cm)

• Drying oven (e.g., Memmert, UF55)

• Ziploc plastic bags (10 × 15 cm)

Prepare microbial inoculum (positive control)

These steps should be performed in a biological safety cabinet using appropriate sterile technique.

• 1In a sterile 100‐ml beaker, reconstitute one vial containing 100 µl A. chroococcum or 100 µl A. brasilense in 20 ml Ashby's mannitol broth or 20 ml DYGS broth, respectively.
• 2Seal the beaker with Parafilm (to minimize the risk of contamination) and incubate 24 hr at 28° ± 2°C with continuous shaking (120 ± 6 rpm).
• 3Add the suspension to 200 ml sterile distilled water in a sterile 500‐ml beaker and stir until the mixture is completely homogenous.
• 4Measure the absorbance at 620 nm using a spectrophotometer and using the appropriate broth as a blank.

  The reading should be between 0.189 and 0.210 for A. chroococcum and between 0.182 and 0.217 for A. brasilense.

• 5Adjust the inoculum to 10⁶ CFU/ml.

  Be sure that all seeds and materials for the bioassay are available and the inoculum is used on the same day that it is prepared. For each microorganism, it is recommended establish a growth curve with a microplate reader or a spectrophotometer.

Treat seed groups

These steps should be performed in a biological safety cabinet using appropriate sterile technique.

• 6Select and count a minimum of 90 soybean seeds of approximately the same size and without stains. Divide them into three equal groups.
• 7Using 250‐ml beakers, immerse one group of seeds in sterile distilled water (the negative control), one in the inoculum from step 5 (positive control), and one with the biofertilizer to be tested. Allow to sit for 1 hr.
• 8Remove seeds with sterile forceps and dry them on sterile absorbent paper.

Sow and inoculate seeds

From this step on, the experiment is carried out in a bioassay room with controlled temperature (average 25° ± 5°C) and relative humidity (49%‐70%).

• 9For each seed, prepare a 9‐oz plastic cup with ∼210 ± 10 g autoclaved soil. Moisten with sterile distilled water at 50%‐60% of moisture retention capacity.

  Moisten the soil to a field capacity of 50%‐60% by weight.

• 10Sow one seed in each cup. Arrange the cups into three replicates per treatment, each with ten experimental units (cups). Allow seeds to germinate.
• 11
  On the 8th day after sowing:

  1. Inoculate each positive control with 2 ml freshly prepared inoculum, each test sample with biofertilizer, and each negative control with distilled water.
     We recommend performing this inoculation in the morning. Be sure to use a proper micropipette with tightly fitting tips and to change tips between treatments.
  2. Add 2 ml Hoagland's nutrient solution without nitrogen to the biofertilizer samples and positive controls.
     We recommend performing this step in the afternoon. We also recommend irrigating on demand (60% of moisture retention capacity) for the duration of the experiment (15 days), removing excess water and allowing gravitational water to drain from the soil.

• 12On day 15 post‐incubation, harvest plants and remove soil from the roots with water.

  Be careful not to damage the roots when removing plants and removing excess water. Washing the roots can be accomplished under a hose in just 30 s. Run the water over a fine‐mesh sieve (0.2‐1 mm) to remove fine heavy particles such as sand, and rinse in water containers to remove coarse heavy particles. We recommend using Wypall disposable wipes to dry the plants after washing.

• 13Measure stem length (SL) using a flexometer to measure the stem with its foliage, from the base of the stem to the apex of the plant.
• 14Measure leaf area (LA) by cutting the cotyledons (the first leaves of the plant) and measuring the leaf area of each plant.

  LA can be calculated with the Montgomery method (MP) by multiplying the length of the blade × the maximum width × the factor 0.75 (Montgomery, 1911). Another method is to use a Portable Leaf Meter or use a digital camera to capture an image of a plant and use special software to analyze the surface area of the leaves. We recommend performing this step on the same day as plant harvest.

• 15To measure dry weight (DW), place the entire plant (stem and foliage) in a paper bag and dry at 60° ± 2°C for 3‐5 days. Let cool in a dry environment (e.g., a Ziploc bag) and then weigh on an analytical balance.

  This should be done in a short time period because the plant tissue will take up water. Avoid formation of a humid environment during cooling. Monitor the moisture content during drying, and stop the process when the sample has reached a consistent weight and moisture content of ∼2% to 5%. Drying can also be done at 60°‐75°C, which will take longer but can have the advantage of preventing roots from being pulverized.

Basic Protocol 2. EVALUATING THE BIOLOGICAL EFFECT OF BIOFERTILIZER BASED ON PGPB UNDER GREENHOUSE CONDITIONS

The genus Bacillus is made up of exo‐ and endophytic bacteria with characteristics that promote plant growth by providing benefits such as solubilization of soil phosphorus, contribution to nitrogen fixation, and production of siderophores (Castaldi et al., 2021). Growth promotion by Bacillus sp. is plant specific, being limited to grasses, legumes, and several vegetables.

Trials in greenhouses using semi‐controlled conditions can demonstrate that microorganisms isolated in the laboratory are effective under normal crop conditions with environmental fluctuations. The number of repetitions and replications over time must ensure confidence and consistency of results. When designing the experiment, the conditions of the test must provide statistically representative results.

This protocol uses broccoli (Brassica oleracea) as a model plant to determine the effectiveness of inoculants based on PGPB (i.e., Bacillus sp.) through the determination of agronomic parameters. It is done via a bioassay under semi‐controlled greenhouse conditions. It uses three treatments: a negative control, a positive control (reference strain), and the biological inoculant to be evaluated. Each treatment has three replicates with ten experimental units (see Statistical Analysis). In planning, it is important to determine the number of treatments, samples, and replications; the experimental design; and the manpower and space availability in the greenhouse. The benefit of this approach is that it provides results in 30 days and can be used for several phases including strain selection and formulation as well as a final quality control parameter.

Additional Materials (also see Basic Protocol 1)

• Bacillus amyloliquefaciens strain Bs006 (Germplasm Bank of Microorganisms‐Agrosavia; National Collections Registry RNC129)

• LB broth (see recipe)

• Broccoli seeds (Brassica oleraceae, e.g., variety Calabrese)

• Autoclaved peat PROMIX PGX

• Fertilizer (NPK 1‐3‐1, e.g., Irricol)

• 50‐well seedbeds (e.g., A&P, ABA50SP)

• Thermometer, hygrometer, and/or greenhouse monitoring sensor

• 1Prepare a 10⁶ CFU/ml inoculum of B. amyloliquefaciens strain Bs006 in 20 ml LB broth as described (see Basic Protocol 1, steps 1‐5).

  The spectrophotometer reading at step 4 should be between 0.189 and 0.210.

Sow and inoculate seed groups

From this step onward, the experiment is carried out in a greenhouse.

• 2Add 100 ± 10 g autoclaved peat PROMIX PGX to each cavity of nine 50‐well seedbeds (three per treatment).
• 3Moisten the soil to a field capacity of 50%‐60% by weight with sterile distilled water.
• 4Using sterile forceps, sow two broccoli seeds in each cavity of the seedbeds.
• 5Add 5 ml positive control inoculum to three seedbeds, 5 ml biofertilizer to three seedbeds, and 5 ml sterile water to the negative control seedbeds.

  Be sure to use a proper micropipette with tightly fitting tips and to change tips between treatments.

• 6Arrange seedbeds in the greenhouse according to the experimental design (see Statistical Analysis). Monitor the temperature and relative humidity for 30 days. Irrigate all cavities with water when necessary.

  We recommend a temperature of 25° ± 5°C and relative humidity of 50%‐70%. You can use a thermometer, hygrometer, or a greenhouse monitoring sensor.

  We recommend irrigating on demand (60% of moisture retention capacity) for the duration of the experiment (30 days).

• 7On day 7 after sowing, select the healthiest and most vigorous plants in each cavity of the seedbed and apply the same amount (5 ml) of each treatment.
• 8On day 14, fertilize each experimental unit with 5 ml fertilizer (NPK 1‐3‐1).

  When mixing or applying fertilizer, follow the directions for use on the label carefully. Mix only the amount that you need for each application. Do not prepare larger amounts to store for future use, as the fertilizer will degrade and become ineffective over time.

• 9On day 21, apply another 5 ml of each treatment.

  We recommend performing this inoculation in the afternoon.

• 10On day 30, harvest plants, remove soil from the roots, and measure SL, DW, and LA as described (see Basic Protocol 1, steps 13‐15). When measuring SL, use a ruler or flexometer.

Basic Protocol 3. EVALUATING THE BIOLOGICAL EFFECT OF BIOFERTILIZER BASED ON SYMBIOTIC NITROGEN‐FIXING BACTERIA

Common beans and other legume plants can undergo symbiosis with PGPB species and may form nodules (Almeida et al., 2022), and this relationship can be utilized to provide nitrogen in farming systems. In this symbiotic relationship, legumes provide rhizobia with nutrients and rhizobia fix atmospheric gas by reduction into ammonia, which is then delivered to the legume inside the root nodule (Hasan et al., 2022). Such symbiotic relationships can fix up to 80% of biologically fixed nitrogen in agricultural fields (Almeida et al., 2022).

Selecting the appropriate range of plant systems is a critical decision. To evaluate strain nodulation effectiveness under controlled conditions, the legume needs to grow for seven weeks or more to exhaust cotyledon nitrogen before the expression of symbiosis becomes evident (Howieson & Dilworth, 2016). Evaluation of effectiveness under controlled conditions is important. Another critical decision is selecting the agronomic variables, such as successful and functional nodules. For this type of analysis, one technique is to count nodules throughout the root, then make a longitudinal cut in the nodules and visually evaluate whether they have a pink color (which indicates the presence of leghemoglobin activity) and determine the weight of the root (Hernández Forte & Nápoles García, 2017). For screening of N₂‐fixing microorganisms, the dry weight of plants is also an important variable because it is related to plant biomass accumulation (Howeison, 2016).

Biological inoculants such as Rhizobium sp. and Bradyrhizobium sp. must demonstrate not only their concentration and purity (by legislation of each country) but also their infectivity and effectiveness in their symbiotic association with legumes. Burton's method involves determining the percentage of plants nodulated (PPN), which provides an evaluation of the efficiency of the biological inoculant in affecting nodulation (Perrone, 2017). Positive plants have three or more nodules located inside an imaginary cylinder with a central axis in the main root, a diameter of 2.5 cm, and a length of 2.5 cm. The results are expressed as a percentage of satisfactorily nodulated plants (SENASA, 2011) and the desired outcome is 80% positive plants.

In this protocol, nitrogen fixation is determined by the efficiency and number of the nodules (Garrido et al., 2019). The location of nodules in the neck of the primary root is related to the infection that occurs in the first ∼6 hr after infection. A greater presence of nodules within the defined cylinder indicates a greater speed of nodulation by the strain (Perrone, 2017) and indicates a competitive advantage over other rhizobia. Four treatment groups are evaluated: a biofertilizer (sample to be tested), a positive control (reference strain), a negative control (water), and a chemical treatment. After the germinated plants are thinned, the sample and positive control are treated weekly without nitrogen while the chemical group is treated with nitrogen. Each treatment has three blocks and each block has ten experimental units (see Statistical Analysis). The protocol is carried out in two parts, the first a biological safety cabinet in the laboratory and the second in a greenhouse.

Additional Materials (also see Basic Protocol 1)

• Rhizobium leguminosarum (ATCC 10004) or Bradyrhizobium japonicum (ATCC 10324)

• LMA broth (see recipe)

• Bean cowpea seeds (Vigna unguiculata L. Walp)

• Biofertilizer: any prototype or final product based on symbiotic nitrogen‐fixing bacteria

• Autoclaved vermiculite (particle size 1‐5 mm)

• Autoclaved river sand

• Hoagland's Nutrient Solution (see recipe), complete and without nitrogen

• Anchor germination paper #38 (10 × 15 in.)

• Ziploc plastic bags (15 × 20 cm)

• Sterile 250‐ml beakers

• Seedling bags (21 × 12 cm)

• Thermometer, hygrometer, and/or greenhouse monitoring sensor

• 10‐100 ml Dispensette bottle‐top dispenser (Brand, BR4600171‐1EA)

• 1Prepare an inoculum of R. leguminosarum or B. japonicum in 20 ml LMA broth as described (see Basic Protocol 1, steps 1‐5), but stir the inoculum for 72 hr (not 24 hr) and adjust the final inoculum to 10⁸ CFU/ml.

  The spectrophotometer reading at step 4 should be between 0.191 and 0.211.

Prepare seeds by treatment group

These steps should be performed in a biological safety cabinet using appropriate sterile technique.

• 2For each of the four treatment groups, select and count a minimum of 120 bean seeds of approximately the same size and without stains.
• 3Moisten a 10 × 15–in. sheet of germination paper with enough sterile distilled water to double its weight. Spread 30 seeds on the paper and place a damp sheet of paper on top. Fold the bottom 1 in. of the paper up, then roll the paper up and place it in a Ziploc bag. Repeat four times for each treatment group (total 120 seeds per treatment group).
• 4Place bags in a bioassay room at 28° ± 2°C for 48 hr.

  If you do not know how long the seeds will take to germinate, open the test up after one day to see if any seeds have germinated. The seeds must have healthy sprouts and develop a radical ∼1 cm in length.

• 5Using forceps, remove the germinated seedlings for the positive control group and transfer to a 250‐ml beaker containing inoculum from step 1. Repeat to immerse the sample seedlings in biofertilizer and the positive control and chemical treatment seedlings in sterile distilled water. Allow each beaker of seeds to sit for 20 min.

  Handle seedlings carefully to avoid damaging the radicle. When applying commercial biofertilizer, follow the directions for use on the label carefully.

• 6Remove bean seedlings from the sterile distilled water or inoculum with sterile forceps and dry them on sterile absorbent paper.

Sow seeds

From this step on, the experiment is carried out in a greenhouse.

• 7Mix 15 kg autoclaved vermiculite and 7.5 kg autoclaved river sand (2:1 ratio) in a clean plastic bowl.

  All materials must be rhizobia‐free. Unless contaminated by dust, vermiculite is rhizobia free when obtained from the factory, as are the chemicals, new containers, and tap or distilled water. Nonetheless, we recommend three cycles of sterilization at 121°C, 20 PSI for 1 hr.

• 8Moisten vermiculite‐sand with 15 L distilled water.
• 9Add 1 kg moistened vermiculite‐sand mix to a seedling bag, using 30 bags for each treatment (one bag for each experimental unit).
• 10In each bag, make two holes in the vermiculite‐sand mix, each 1 cm deep, and sow one seed in each hole using sterile forceps.
• 11Place bags on a table in the greenhouse, distributing them evenly and randomly. Monitor the temperature and relative humidity for 60 days. Irrigate seedling bags with distilled water when necessary (apply 100 ml distilled water per bag).

  We recommend a temperature of 16.9°‐36.6°C and relative humidity of 11%‐66%. You can use a thermometer, hygrometer, or a greenhouse monitoring sensor. We also recommend irrigating on demand using a Dispensette.

• 12On day 7 after sowing, hand‐thin by selecting the healthiest and most vigorous plant in each bag and removing the other plant.

  Selected plants should measure at least 10 cm tall.

• 13Apply 10 ml Hoagland's nutrient solution to the selected plants 1 day a week for 7 weeks.

  For the positive control or commercial biofertilizer, apply Hoagland's nutrient solution without nitrogen. For chemical treatment, use Hoagland's complete nutrient solution. For the negative control, use only distilled water.

• 14After 60 days, harvest plants and remove the soil from the roots with water.

  Be careful not to damage the roots when removing plants and removing excess water. The process of washing roots can be accomplished under a hose in just 30 s.

• 15Use a ruler or meter to measure root length (RL) and stem length (SL). For RL, measure the distance along the long axis of the root from the apex of the plant. For SL, measure from the base of the stem to the apex of the plant.
• 16
  Measure the percentage of plant nodulation (PPN, %).

  • a.
    Cut the root of the plant using scissors.
  • b.
    Count nodules in the crown and lateral regions within a 2.5 × 2.5–cm cylinder around the center of the root (Fig. 2).
    It is important to know effective and non‐effective nodules by visual observation of the intensity of their pink color. Nodules with pink coloration are considered effective. A plant is considered positive for nodulation when effective nodules are present within the counting area. Assign positive plant nodulation to plants that have three or more nodules located within the cylinder.
  • c.
    Determine effectiveness using the equation:

    $$\mathrm{PPN}\left(\%\right)=\frac{\mathrm{number}\mathrm{of}\mathrm{plants}\mathrm{with}\mathrm{positive}\mathrm{nodulation}}{\mathrm{total}\mathrm{number}\mathrm{of}\mathrm{plants}}\times 100$$

Figure 2.

Cylinder (2.5 × 2.5 cm) used for quantification of nodules by the PPN method (Perrone, 2017).

• 17Measure DW of the entire plant (stem, foliage, and roots with nodules) as described (see Basic Protocol 1, step 16).

Basic Protocol 4. EVALUATING THE BIOLOGICAL EFFECT OF BIOFERTILIZER BASED ON AMF

AMF are obligate biotrophs, which colonize the roots of 72%‐80% of terrestrial plant species to establish symbioses. AMF enhance the uptake of essential nutrients including phosphorus, zinc, and nitrogen. They may also increase plant resistance toward pathogens and other abiotic stresses such as drought and salinity (Salomon et al., 2022). Bioassays with AMF are performed under controlled or semi‐controlled conditions. The bioassay is designed to control the minimum requirements of commercial inoculants that contain viable propagules and colonize the roots of host plants with a sterile substrate. It is important to carry out experiments in host plants that are sensitive to mycorrhizae (e.g., corn, sorghum, kudzu, and onion), with a positive growth response and a significant colonization of at least 20% (Salomon et al., 2022). Host species such as onion and kudzu are used to preserve and multiply the AMF inoculum due to their known affinity (Criollo et al., 2021).

The following protocol measures the percentage of mycorrhizal colonization in a host plant after a limited time, so that only primary infections occur. At the end of the bioassay (∼4 months), the percentage of infection is determined by examining stained root samples on slides. The total number of fields (intersection between the root and a horizontal transect) and the number of mycorrhized fields are determined. Mycorrhization is established by the presence of structures characteristic of AMF within the root of the plant, i.e., arbuscules, internal mycelium, or vesicles. Inoculum viability and biological activity are the only outcomes of this measurement, and the results should not be represented otherwise.

This protocol is carried out in two parts: root colonization by AMF, which is carried out in a greenhouse, and staining of colonized roots, which is conducted in the laboratory. In the laboratory, all solutions and equipment coming into contact with roots must be sterile. The bioassay has a randomized block experimental design, with four blocks per treatment and five experimental units per block. An experimental unit corresponds to an individual plant; during sowing, a block corresponds to five cavities for each treatment. Each treatment (the biofertilizer to be analyzed, a positive control, and a negative control) thus requires four seedbeds with five cavities. It is essential to follow this experimental design.

It should be noted that standardization of experimental conditions does not mean standardization of the environmental conditions to which fungi will respond. Every fungus in an inoculum can perceive the same host in different ways and react differently to the same medium. Consequently, it is essential to take into account any disparities between the conditions of the assay and the conditions at the site(s) where the test fungi were found.

Additional Materials (also see Basic Protocol 1)

• Autoclaved quartzite sand

• Biofertilizer: any AMF‐based prototype or final product

• Claroideoglomus etunicatum strain Gl07 (Germplasm Bank of Microorganisms‐Agrosavia, National Collections Registry RNC129)

• Rhizoglomus irregularis strain Gl08 (Germplasm Bank of Microorganisms‐Agrosavia; National Collections Registry RNC129)

• Seeds: kudzu (Pueraria thomsonii, e.g., variety Montana) or scallion (Allium fistulosum, e.g., variety Nebuka)

• Hoagland's nutrient solution (see recipe), complete

• 10% KOH (see recipe)

• 10% H₂O₂ (see recipe)

• 1 N HCl (see recipe)

• 0.05% trypan blue (see recipe)

• Lactoglycerol (see recipe)

• Sterile plastic bowl

• 72‐cavity seedbed, plastic (e.g., A&P, A‐BA72)

• Seedling bags (21 × 12 cm)

• Scalpel (no. 4) and surgical blades (no. 20; Paramount, 2460)

• 50‐ml centrifuge tubes (Falcon, 352070)

• 80° ± 2°C water bath (e.g., Memmert, WNB 7)

• Petri dishes, 150 × 25 mm (Nest, 715001)

• Glass slides (Brand, 474701)

• Glass cover slips (Brand, 470045)

• Microscope (e.g., Olympus, CX31RBSFA) with 40× objective

• 1Mix autoclaved soil and quartzite sand 1:1 by volume in a clean and disinfected plastic bowl.
• 2For each treatment, set up four seedbeds with five cavities each. Label each seedbed with treatment, date, and type of seed.
• 3Add 100 g soil‐sand mix to each seedbed cavity for all treatments.
• 4Add 1 ± 0.01 g biofertilizer to be tested to the appropriate seedbeds and 1 ± 0.01 g reference material for the positive control.

  Kudzu seeds are inoculated with R. irregularis and scallion seeds with C. etunicatum. The reference materials contain 80 spores/g.

• 5Sow two kudzu or long scallion seeds per cavity in all seedbed cavities for all three treatments.
• 6Cover the cavities with an additional 20 g soil‐sand mix.
• 7Arrange seedbeds in the greenhouse following a random block experimental design. Irrigate each seedbed with 10 ml distilled water every 3 days for 1 week.

  Be sure to use a proper micropipette for irrigation.

• 8
  On the 7th day after sowing:

  1. Hand‐thin the plants by selecting the healthiest and most vigorous plant in each cavity and removing the other plant.
  2. Apply 2 ml Hoagland's complete nutrient solution to the positive control and biofertilizer treatments once a week for 6 weeks. Use distilled water for the negative control.
  3. Continue to irrigate all treatments as in step 8 until the end of the first 7 weeks.
• 9
  In week 8:

  1. Prepare five seedling bags per treatment with ∼1 kg soil‐sand mix.
  2. Transfer one plant from each cavity to one seedling bag. Do this for each treatment.
  3. Apply 10 ml Hoagland's complete nutrient solution to the positive control and biofertilizer treatments once a week until the end of week 13. For the negative control, use distilled water.
  4. Irrigate all seedling bags with 100 ml distilled water once a week using a dispensette. Stop irrigating 20 days before harvesting the plants.
     Eliminating irrigation prior to harvesting is necessary to stress both the plants and the AMF. This process must potentiate sporulation of the fungus and colonization of the roots.
• 10After 4 months, harvest plants and remove the soil from the roots with water. Store plants individually in plastic bags for up to 8 days at 4°C.

  Be careful not to damage the roots when removing the plants and removing excess water. Washing of the roots can be accomplished under a hose in just 30 s.

Stain colonized roots (laboratory)

An overview of the staining procedure is illustrated in Figure 3.

Figure 3.

Diagram of staining of AMF‐colonized roots.

• 11Sample the roots from all plants and cut into short segments (2‐4 cm). Group roots by treatment and block for processing.

  When sampling roots to measure mycorrhizal colonization, it is important to select finer, more fibrous roots. Older roots, or roots from plants with taproots or other coarse roots, may have some mycorrhizae, but colonization is usually sparse and consists only of hyphae that often are most visible outside the roots. Older roots also tend to resist clearing due to accumulation of tannins and other compounds.

• 12Place roots in a 50‐ml Falcon tube, making sure the sample fits in the lower 50% of the tube.

  For good results, large volumes of roots may need to be subdivided or subsampled. Each tube must be named with experimental unit and treatment.

• 13Add 10% (w/v) KOH to each tube until the roots are covered but the solution does not fill more than half the tube.

  The tissue may be soaked overnight but be sure to replace KOH with fresh KOH before heat treatment.

  CAUTION: Take care to avoid skin contact with KOH. Use gloves and safety glasses.

• 14Place the tube in an 80° ± 2°C water bath for 15 min. If the roots are slim and fragile, reduce the time to 10 min.
• 15Wash roots with water for 1‐2 min.

  Be careful not to damage the roots and remove excess water. There is a possibility of partial or complete disintegration of lateral roots. Care should be taken not to dissolve parts of the ground tissue (the cortex usually goes first).

• 16 Optional: Immerse roots in a 1:1 (v/v) solution of 10% KOH and 10% H₂O₂ for 10 min, then wash with water.

  Use this step only if the roots are thick and have a high tannin content.

• 17Add 1 N HCl to each tube until the roots are covered and incubate for 10 min. Discard the solution into a liquid chemical container and wash roots with water.
• 18Add 0.05% trypan blue until the roots are covered and place tube in the 80°C water bath for 10 min. Discard the stain into a liquid chemical container and wash roots with water for at least 1 min.

  If you want to leave the roots to be processed the next day, add deionized water and leave covered for 12 hr to remove excess dye.

  CAUTION: Wear gloves and avoid skin contact when handling trypan blue solution.

• 19Place roots in a Petri dish and cut ten 2‐cm pieces per plant (200 pieces per treatment group).
• 20Arrange individual root segments on glass microscope slides (several segments from each sample can fit in parallel). Add a drop of lactoglycerol and apply a cover slip.

  To improve visualization under the microscope, the roots can be pressed gently onto the slide using a cover slip to flatten them.

• 21Assess the presence of visible AMF structures (e.g., arbuscules, hyphae, and vesicles; Fig. 4) under a microscope at 400× magnification (10× ocular/40× objective). Count at least ten intersections for each piece of root for a total of 100 intersections.

  Observations must be taken in triplicate. This method allows for a simple analysis of colonization at each intersection of the eyepiece crosshair.

Figure 4.

Kudzu root staining showing AMF structures (arbuscules, vesicles, intraradical hyphae, and spores). Bright‐field image taken at 40× magnification.

• 22
  Calculate the percentage of colonization using the equation:

  $$\mathrm{Colonization}\left(\%\right)=\frac{\mathrm{number}\mathrm{of}\mathrm{intersections}\mathrm{on}\mathrm{AMF}}{\mathrm{number}\mathrm{of}\mathrm{intersections}\mathrm{on}\mathrm{stained}\mathrm{root}}\times 100$$
• 23
  Calculate the effectiveness of each treatment (commercial biofertilizer and positive control) using the equation:

  $$\mathrm{Effectiveness}\left(\%\right)=\frac{\mathrm{plants}\mathrm{colonizated}\mathrm{by}\mathrm{AMF}}{\mathrm{total}\mathrm{plants}\mathrm{inoculated}\mathrm{with}\mathrm{AMF}}\times 100$$

REAGENTS AND SOLUTIONS

Ashby's mannitol broth

• In a glass bottle (e.g., Schott bottle), combine:

• 10 g mannitol (Merck, 443907)

• 0.2 g KH₂PO₄ (Sigma‐Aldrich, P0662)

• 0.2 g MgSO₄·7H₂O (Merck, 1374361)

• 0.2 NaCl (Merck, 106400)

• 0.2 g CaSO₄ (Merck, 255696)

• 5 g CaCO₃ (Merck, C4830)

• 1000 ml distilled water

• Adjust to pH 7.2 using HCl or NaOH

• Sterilize by autoclaving 15 min at 120°C

• Store up to 1 month at 4°‐8°C

DYGS broth

• In a glass bottle (e.g., Schott bottle), combine:

• 0.5 g K₂HPO₄ (Merck, 1551128)

• 0.5 g MgSO₄·7H₂O (Merck, 1374361)

• 2 g glucose (Merck, G8270)

• 1.5 peptone (Merck, 68971)

• 2 g yeast extract (Merck, 1.13885)

• 1.5 g glutamic acid (Merck, G0355000)

• 2 g malic acid (Merck, PHR1273)

• 1000 ml distilled water

• Adjust to pH 6.5‐6.7 using HCl or NaOH

• Sterilize by autoclaving 15 min at 120°C

• Store up to 1 month at 4°‐8°C

Hoagland's nutrient solution

For Hoagland's complete nutrient solution, add 1 ml Hoagland's micronutrient solution (see recipe) to 999 ml Hoagland's macronutrient solution (see recipe) in a 1000‐ml volumetric flask.

For Hoagland's nutrient solution without nitrogen, omit solutions that contain nitrogen ((NH₄)H₂PO₄ and NaNO₃).

Hoagland's macronutrient solution

• In a 1‐L volumetric flask, combine:

• 6 ml of 1.0 M K₂HPO₄ (final 6 mM)

• 4 ml of 1.0 M NaNO₃ (final 4 mM)

• 2 ml of 1.0 M (NH₄)H₂PO₄ (final 2 mM)

• 1 ml of 1.0 M MgSO₄·7H₂O (final 1 mM)

• Distilled water to a final volume of 1 L

• Store up to 1 month at 4°‐8°C

Hoagland's micronutrient solution

• In a 1‐L volumetric flask, combine:

• 6 ml of 1.864 g/L KCl

• 4 ml of 0.773 g/L H₃BO₃

• 2 ml of 0.169 g/L MnSO₄

• 1 ml of 0.288 g/L ZnSO₄·7H₂O

• 1 ml of 0.062 g/L FeSO₄·7H₂O

• 1 ml of 0.04 g/L Na₂MoO₄·2H₂O

• 1 ml of a solution containing 1.118 g/L titriplex III and 0.834 g/L iron sulfate (NaFeEDTA, 10%)

• Distilled water to a final volume of 1 L

• Store up to 1 month at 4°‐8°C

Hydrochloric acid (HCl), 1 N

Add 86 ml of 37% HCl (Merck, 109057) to 1000 ml distilled water. Mix well and store up to 1 month at 4°‐8°C.

Hydrogen peroxide (H₂O₂), 10%

Dilute 30% H₂O₂ (Merck, 107209) to 10% in distilled water. Mix well and store up to 1 month at 4°‐8°C.

Lactoglycerol

• Prepare in an amber glass container:

• 125 ml lactic acid (Merck, 100366)

• 250 ml glycerol (Merck, 104091; 85% final)

• 125 ml distilled water

• Mix with frequent agitation until homogeneous

• Store up to 1 month at 4°‐8°C

LMA broth

• In a glass bottle (e.g., Schott bottle), combine:

• 10 g mannitol (Merck, 443907)

• 0.5 g yeast extract (Merck, 1.13885)

• 0.5 g K₂HPO₄ (Merck, 1551128)

• 0.1 g MgSO₄·7H₂O (Merck, 1374361)

• 0. 2 g NaCl (Merck, 106400)

• 18 g agar (Merck, 05039)

• 1000 ml distilled water

• Adjust to pH 6.8 using HCl or NaOH

• Sterilize by autoclaving 15 min at 120°C

• Store up to 1 month at 4°‐8°C

Luria Bertani (LB) broth

• In a glass bottle (e.g., Schott bottle), combine:

• 10 g tryptone (Merck, T7293)

• 5 g yeast extract (Merck, 1.13885)

• 10 g NaCl (Merck, 106400)

• 20 g agar (Merck, 05039)

• 1000 ml distilled water

• Adjust pH to 6.8‐7.2 using HCl or NaOH

• Sterilize by autoclaving 15 min at 120°C

• Store up to 1 month at 4°‐8°C

Potassium hydroxide (KOH), 10% (w/v)

Add 100 g KOH (Merck, 105033) to 1 L distilled water. Mix well and store up to 1 month at 4°‐8°C.

Trypan blue, 0.05%

Prepare 500 ml lactoglycerol (see recipe) with 0.25 g trypan blue (Merck, 111732). Mix until homogeneous and store up to 1 month at 4°‐8°C.

CAUTION: Take care to avoid breathing dust or getting any in your eyes when handling trypan blue powder. Wear gloves to protect your hands when using dye powders or solutions.

COMMENTARY

Background Information

The sustainable production of food for a population that is predicted to reach 9.7 billion people by 2050 is one of the great issues of the 21st century. Over the past two centuries, techniques for increasing food production yields have relied mainly on chemical pesticides and mineral fertilizers (Salomon et al., 2022). However, the manufacturing of these chemical fertilizers involves some of the most energy‐intensive processes in the world and is often dependent on limited resources, as in the case of phosphorus fertilizers. Moreover, some crops have low phosphorus fertilizer use efficiencies, which leads to low fertilizer uptake by plants. According to Walling and Vaneeckhaute (2020), the intensive use of fertilizers in food production systems is a significant contributor to agricultural world greenhouse gas emissions and can have adverse effects on biodiversity and environmental sustainability. Without dietary modifications or a decrease in food waste, projections show that present yield trends would not be sufficient to supply the demand for food in the next decades (Daniel et al., 2022). Other urgent problems include the appearance of novel crop pests, the development of pesticide resistance, the rising demand for pesticide‐free food, and the improvement of food safety (Daniel et al., 2022). Thus, there is a quickly growing interest in reducing our dependency on agrochemicals through the use of bioproduct‐based inoculants such as microbial inoculants. Commercial microbial biofertilizers include wildly popular rhizobia products as well as products based on other species such as Bacillus sp. or Trichoderma sp. that improve plant vigor and have the potential to reduce the demand and use of fertilizer chemicals (Fasusi et al., 2021). Growing scientific evidence demonstrating numerous advantages that biofertilizers bring to crops in terms of growth and yield has attracted the attention of end users (Basiru et al., 2021).

Microbial biofertilizer use has an annual growth rate of 10%‐12% in the market. By the year 2016, the market size of the global biofertilizer market is projected to grow at a rate of 14.2% to reach USD 3.1 billion (Joshi & Gauraha, 2022). Different interactions between plant roots and soil microorganisms have an impact on plant nutrition, either directly by affecting the availability of minerals or indirectly by improving absorption efficiency through the stimulation of plant root growth. In general, the success of biofertilizers, especially those based on PGPB and AMF, requires research on screening, mass production, formulation, and quality control. In particular, quality control is crucial to improving the education and business management of this type of bioproduct. Hence, there is an urgent requirement to adopt universal, highly reproducible methods to evaluate the quality of commercial biofertilizers in order to deliver a consistent product to field crops and greenhouse conditions. This will also help strengthen regulatory mechanisms for microbial biofertilizers. To manage the microbiological products in support of clients, quality supervision and final quality control of product is extremely important and should be consistently attained (Agarwal et al., 2021). The criteria used to determine quality are limited to the viability and conservation of specific bacteria or fungi, but the most important, and sometimes difficult to assess, is biological activity.

The manufacture of high‐quality inoculants increases the potential to promote plant growth in inoculated seeds or plants, and the assessment of biofertilizer quality has long been a topic of study. For example, large‐scale production of inoculants is associated with many technical difficulties. For biofertilizers based on bacteria, media and growth conditions (temperature, pH, time) are critical for maintaining good physiological conditions. For biofertilizers based on AMF, strain‐host specificity is an important factor in choosing hosts for the multiplicity of AMF strains. In all cases, the availability of competent and well‐trained operators is crucial to ensuring that the right methodologies are implemented. In addition, minimizing production costs and preserving the pure microbial culture throughout the process are important factors. As a result, the product quality will be improved (Mishra & Barolia, 2020).

The goals of quality control methods for biofertilizers are to determine and count the microorganisms present in a given formulation at a particular moment and to measure indicators that the quality will be preserved over the storage time. The main activities and parameters are monitoring batch variability, recovery, purity, and enumeration of cells; positively identifying microorganisms from the carrier; measuring moisture content and other conditions that support growth and survival in the formulation; and evaluating products from various supply chains (Yadav & Chandra, 2014). The efficiency of a biofertilizer must be evaluated under laboratory or greenhouse conditions, and thus bioassays must be repeatable and reliable to determine accurate effectiveness. Variability in an assay's components may be a source of inconsistent results, but few standardized bioassays are available. Few papers describe the methodology used to develop bioassays for a specific purpose, and it is difficult to design bioassay procedures suitable for evaluating a microorganism‐plant relationship without these details.

Standardization for measuring efficacy has been developed for a few biofertilizers based on Rhizobium sp., for example, but not for other biofertilizers (Yadav et al., 2020). In general, standardization of biological activity allows scientists and regulators to compare products. In addition, several authors and laboratories have expressed a desire for commonly accepted standard methods and criteria in other bioproducts. The protocols presented here demonstrate the capacity to assess different types of biofertilizers based on bacteria or fungi as an active principle.

Basic Protocols 1 and 2 focus on biofertilizers based on PGPB such as Azospirillum sp. (A. brasilense, A. lipoferum, A. amazonense, A. halopraeferens, and A. irakense), Acetobacter diazotrophicus, Herbaspirillum sp., Bacillus sp., and Pseudomonas sp. under laboratory or greenhouse conditions and have the advantage of minimizing both cost and time. Basic Protocol 3 focuses on biofertilizers based on symbiotic nitrogen‐fixing bacteria such as Rhizobium sp., but can be extended to other microorganisms such as Bradyrhizobium sp., Sinorhizobium sp., Azorhizobium sp., Mesorhizobium sp., and Allorhizobium. In the case of biofertilizers based on AMF, it is critical to choose the perfect plant host using a model plant, an option that is provided in Basic Protocol 4. These protocols can be extended to other microorganisms that improve plant growth and can reproduce to reduce variability. All protocols can be used in multiple stages of biofertilizer development, including selection of the microorganism, characterization of the microorganism, formulation compatibility, and in vitro evaluation of effectiveness.

Critical Parameters

For a well‐established effectiveness determination, the experimental design must be statistically robust. For Basic Protocol 1, experiments must be arranged in a completely randomized design with a minimum of ten experimental units for each replicate, with a minimum of three replicates per treatment. For Basic Protocols 2–4, a randomized block design is often useful. Using this design, treatments are allocated into replicates, which are sown in blocks, and the blocks are arranged such that any evident variation in the greenhouse is exposed to a discrete replicate.

In Basic Protocols 3 and 4, it is important to assess the effectiveness under controlled conditions using the following recommendations and treatments. (1) Except for the nitrogen in the seed, no other mineral nitrogen that is accessible to plants should enter the system. This makes it possible to track N₂ fixation as a direct function in the plant. (2) Controls must be included in the treatments. It is important to include a reference strain and chemical control treatments. The chemical control treatment must have N available at a non‐limiting rate, such that growth can be compared with the commercial biofertilizer and reference strain treatments, and relative effectiveness of strains can be assessed by comparison.

Some additional recommendations are listed below.

Seed quality

For biological activity, seed quality is critical. This is defined as the degree to which a seed lot meets standards that determine the seed quality (genetic, physical, physiological, and health parameters). The attributes correspond to genetic purity, physical purity, and germination capacity (an indication of the proportion of live seeds that can produce normal seedlings). Other attributes include moisture content, seed vigor, and health. Seed quality must fulfill all these attributes. For all protocols, it is important to ensure that the seed to be used has high quality, especially a high percentage of germination, high genetic and physical purity, and an absence of disease (ISTA, 2016).

Seed germination percentage

The percentage of seed germination is assessed by the International Rules for the Analysis of Seeds found in the ISTA Manual (ISTA, 2016). The test is based on 400 seeds, which can be divided into four repetitions of 100 seeds. It is important that seeds be separated on the substrate to prevent contact between seedlings before counting and removal. The method is carried out in the laboratory under controlled conditions in order to ensure consistent, fast, and complete germination. Conditions are described in the ISTA Manual and include optimal humidity. Additionally, conditions are normalized, so results can be reproduced within limits on variation of the random sampling (ISTA, 2016). The germination rate should be at least 75% for Basic Protocol 1 and 80% for Basic Protocols 2–4.

Purity of microbial inoculum

Microbial purity is an important quality control parameter due to the competition that may occur with contaminants for nutrients and space, producing metabolites as a defense. Evaluation of biological activity relies on defining the efficacy of the biofertilizer and the biological stability of the active ingredient (Hermann et al., 2015). Contamination should be avoided by using strict aseptic technique in a circulating airflow cabinet (class II) and by following all safety guides.

Reference strains

It is important to use biochemically or molecularly characterized or certified reference strains. The reference strain should be acquired from a recognized culture collection such as the ATTC. It is important to construct a calibration curve for each reference strain by diluting the original culture with growth medium to produce at least five points and determine a plate count at each dilution.

Substrate type

In greenhouses, different organic support substrates are used that can positively or negatively affect growth outcomes. Substrates used in all protocols must have the correct chemical, physical, and microbiological characteristics. A good substrate must enable adequate transfer of nutrients to the roots. Physical characteristics such as texture, granulometry, and porosity must be considered, so that root asphyxia does not occur. Another important factor is contaminating microbial load. Substrates for all of these protocols must be sterile. Substrates such as vermiculite, soil, and river sand must be autoclaved three times for 1 hr, unlike peat, which can be purchased sterile for immediate use (Carrasco et al., 2005).

Substrate moisture

Substrates such as soil supply and retain nutrients and moisture (Piñón‐Villarreal et al., 2013), and thus require less water and nutrients to be provided. Substrates such as vermiculite and sand need more frequent application of water and nutrients (between 20% and 30%; Papadopoulos et al., 1992), and there is more accumulation of moisture and nutrients in the medium due to the type of substrate (Lopez‐Iglesias et al., 2014). It is important to irrigate on demand for all protocols. It is recommended to perform a preliminary assessment of the substrate that most suits the needs of the biological material to ensure that plant growth is optimal.

Re‐inoculation

For Basic Protocol 3, the beginning of symbiosis may be affected by the size of the seeds, because larger seeds have greater energy reserves and a longer time to cotyledon senescence, which can improve symbiosis (López et al., 2022). To ensure symbiosis, it is important to perform a second inoculation when the tap and secondary roots are already developed.

Equipment

It is necessary to carry out processes using calibrated and verified laboratory equipment to obtain reliable, precise, and reproducible results. All equipment should be calibrated and inspected within the manufacturer‐specified intervals to ensure the quality of the data.

Temperature and humidity control in the greenhouse

Climate control inside a greenhouse is the most important factor. If greenhouse conditions are not favorable, the plants will be affected. During the day, plants open their stomata to favor gas exchange and transpiration. If the relative humidity drops or the temperature rises, the plant will close its stomata, transpiration will decrease, and photosynthetic production will decrease. Thus, all protocols in a greenhouse must be performed with temperature and humidity control.

Troubleshooting

When a problem is encountered with a protocol, every step should be reviewed. A few common problems and solutions are listed in Table 1.

Table 1.

Troubleshooting for High‐Throughput Laboratory or Greenhouse Evaluation of the Biological Efficacy of Biofertilizers

Problem	Possible cause	Solution
Inoculum contamination	Cross‐contamination or mishandling of materials	Prepare new inoculum; ensure microbiological control (sterilization) of surfaces and environments
Low seed germination (%)	Poor seed quality	Change seed lot
Bacterial or fungal contamination of substrate	Poor sterilization and/or aeration	Sterilize substrates and allow them to air. Perform 1‐hr autoclave cycles every 24 hr to ensure cooling and aeration between cycles.
High humidity in substrate	Excessive irrigation	Pause watering
Symptoms of disease or pests in plants	Seed‐associated diseases or inadequate bioassay monitoring	Perform new assembly
No root penetration by dye	Thick and lignified roots	Leave the roots in dye for up to 24 hr; after applying 10% KOH, immerse roots in 1:1 (v/v) 10% KOH and 10% H2O2 for 10 min
	Dye expired or prepared poorly	Prepare new dye

Statistical Analysis

Data can be processed and analyzed by descriptive and inferential statistics such as means, frequency distributions, and percentage using statistical software. Using the Shapiro‐Wilk and Bartlett tests, respectively, the normality and homogeneity of variance of the data must be verified (ɑ = 0.05). The significan effects of treatments on measured variables can be determined using a one‐way analysis of variance (ANOVA) followed by a post hoc test such as Duncan's or Tukey's test. The Pearson's correlation coefficient can be used to determine the relationship between plant growth (variable response) and some nitrogen fixation parameters such as % nodulation (Basic Protocol 3) or effectiveness (Basic Protocol 4).

Understanding Results

As an example, we describe how to understand the results from an assessment of the effectiveness of two AMF strains (Basic Protocol 4). The same analysis can be done with data obtained from Basic Protocols 1–3. In this example, we evaluated the effectiveness of R. irregularis (strain Gl08) and C. etunicatum (strain Gl07) using kudzu and scallion as model plants in a greenhouse experiment comprised of four blocks, each with five experimental units per treatment. After 45 days, colonization (%) and effectiveness (%) were determined. In scallion, colonization was higher for R. irregularis (9.9%) than C. etunicatum (6.6%). In scallion, colonization was higher for C. etunicatum (22.0%) than R. irregularis (8.2%). The control treatment had 0% colonization, suggesting a good execution of the protocol and confirming that the substrate had no contaminating AMF that could interfere with the results (Table 2).

Table 2.

Colonization Rates (%) of R. irregularis (strain Gl08) and C. etunicatum (strain Gl07) in Kudzu and Scallion

	Colonization (%) in scallion		Colonization (%) in kudzu
Block	C. etunicatum	R. irregularis	C. etunicatum	R. irregularis	Control treatment a
1	0.2	5.8	19.0	0.0	0.0
2	0.0	20.6	30.2	2.8	0.0
3	12.6	7.6	30.0	30.0	0.0
4	13.4	5.6	8.6	0.0	0.0
Average	6.6	9.9	21.9	8.2	0.0

^a Corresponds to combination of all negative controls.

In kudzu, effectiveness was higher for treatment with R. irregularis than treatment with C. etunicatum (70% vs. 25%; Fig. 5). Mycorrhizal inoculation with R. irregularis had a positive effect on kudzu biomass production (p ≤ .05 versus control). The scallion group inoculated with R. irregularis showed the highest concentrations of mycorrhized plants (p ≤ 0.05) with respect to the rest of the treatments in this model plant. This result shows efficient mycorrhizal function and an affinity for the Glomeraceae family of AMF in kudzu (Senés‐Guerrero et al., 2014).

Figure 5.

Effectiveness (%) of R. irregularis (strain Gl08) and C. etunicatum (strain Gl07) using kudzu and scallion as model plants. Data are mean (n = 15); treatments with different letters (a,b) are significantly different (p ≤ 0.05) according to the Tukey test (95%).

Figure 5 shows an example of how effectiveness is plotted. In scallion, the affinity was higher with C. etunicatum (effectiveness = 68.0%) and lowest with R. irregularis (15.0%). There were also significant differences (p < .05) among strains and model plants. Based on these results, and with other variables such as the plant weight, this study demonstrates that effectiveness fits with growth parameters and is significantly influenced by the presence of mycorrhizaed plants. The difference in mycorrhization frequency between plants and strains is probably due to soil composition, environmental conditions, and the relation between plant hosts, and for these reasons the use of a commercial biofertilizer must be optimized to select the best host plants in sterilized substrates under controlled conditions. According to Salomon et al. (2022), the desired outcome at the end of the bioassay is a positive growth response and a significant AMF effectiveness of at least 20%, which can be established by following Basic Protocol 4.

Time Considerations

Evaluation of biological effectiveness of growth‐promoting bacteria takes 20 days in the laboratory and ∼30‐35 days in the greenhouse. Evaluation of symbiotic nitrogen‐fixing bacteria in the greenhouse takes 2 months. Evaluation of AMF takes ∼40‐50 days, depending on the analysis of the percentage of mycorrhization. These times depend on incubation times for the reference strains, drying of plant materials, and data collection.

Author Contributions

Ivonn Gelvez‐Pardo: data curation, formal analysis, investigation, methodology, writing (original draft, review and editing); Laura Lobo‐Berbesi: data curation, formal analysis, investigation, methodology, writing (original draft, review and editing); Adriana Marcela Santos‐Diaz: conceptualization, data curation, formal analysis, investigation, methodology, writing (original draft, review and editing).

Conflict of Interest

The authors declare that they have no conflict of interest.

Gelvez‐Pardo, I. , Lobo‐Berbesi, L. , & Santos‐Díaz, A. (2023). Biological efficacy of plant growth–promoting bacteria and arbuscular mycorrhizae fungi: Assessments in laboratory and greenhouse conditions. Current Protocols, 3, e732. doi: 10.1002/cpz1.732

Data Availability Statement

Data that support the findings of this study are available from the corresponding author upon reasonable request.