- Open Access
Loss of arbuscular mycorrhizal fungal diversity in trap cultures during long-term subculturing
IMA Fungus volume 4, pages 161–167 (2013)
Long-term successional dynamics of an inoculum of arbuscular mycorrhizal fungi (AMF) associated with the maize rhizosphere (from traditionally managed agroecosystems in Los Tuxtlas, Veracruz, Mexico), was followed in Bracchiaria comata trap cultures for almost eight years. The results indicate that AMF diversity is lost following long-term subculturing of a single plant host species. Only the dominant species, Claroideoglomus etunicatum, persisted in pot cultures after 13 cycles. The absence of other morphotypes was demonstrated by an 18S rDNA survey, which confirmed that the sequences present solely belonged to C. etunicatum. Members of Diversisporales were the first to decrease in diversity, and the most persistent species belonged to Glomerales.
Arbuscular mycorrhizal fungi (AMF) belonging to the phylum Glomeromycota are important soil organisms that form mutualistic associations with plants, and which are involved in the uptake and transport of mineral nutrients to plant roots (Barea et al. 2002). Up to 90 % of analysed plant species are able to form this symbiosis (Smith & Read 1997). Soil structure is improved by the presence of AMF, which likely represents a large component of the microbial biomass (Sieverding & Oehl 2005).
Several studies have suggested that there has been very little selective pressure for either symbiotic partner to develop a high degree of specificity (Harley & Smith 1983, Law & Lewis 1983, Clapp et al. 1995). However, Chanway et al. (1991) noted several factors that may result in strong selection for plant host specificity, such as geographical distance, environmental heterogeneity, host identity, and habitat, which might also affect distribution of AMF. While some authors have noted a lack of host specificity (Merckx et al. 2012), others report that certain hosts show preferences for certain AMF (Yang et al. 2012).
Edaphic conditions and cultural management in different agroecosystems are factors that may influence soil AMF (Oehl et al. 2003, 2004, 2005). Very few studies describe the selective pressure on AMF species when they are taken from their natural environments to trap cultures (Antunes 2012, Oliveira 2010). Cuenca et al. (2003) reported that AMF propagation in trap cultures exhibits difficulties because exact natural conditions cannot be reproduced; this causes a bias towards proliferation of species that are able to tolerate greenhouse conditions, and that associate better with the plant host under the specific trap culture conditions. Leal et al. (2009) have also reported that the trap-plant technique does not necessarily facilitate AMF identification from a specific ecosystem, since diverse factors in the trap cultures might prevent the propagation of all the species present in the initial inoculum. Factors such as the soil or substrate type used and handling, can also include the broad variety of plant species used for propagating AMF in trap cultures (Yao et al. 2010). These results do not necessarily support or contradict the observations that either multiple infections or AMF selectivity occur between certain host-fungus combinations in co-existing plants of a natural community. They do, however, indicate that mycorrhizal fungi respond differentially according to the host species; these differences could thus result in a selective pressure that favours certain host-fungus combinations (Bever 2002, Sanders & Fitter 1992).
This study aims to present a long-term successional analysis of an AMF maize rhizosphere community from traditionally managed sites in Los Tuxtlas, Veracruz, maintained in trap cultures of Bracchiaria comata in continuous subculturing for almost eight years.
Materials and Methods
Sampling was carried out in the Los Tuxtlas, Biosphere Reserve area (UNESCO — MAB Biosphere Directory), in the state of Veracruz, Mexico (18° 24′ 56″ N — 18°26′33″ N; 94° 56′ 53″ W — 94° 58′ 18″ W), situated at an altitude of 238.39 ± 37.45 m (García et al. 2009). The annual precipitation is 2 000–2 500 mm; the weather is characterised as Am (f), i.e. tropical warm and humid, and the mean annual temperature ranges from 22 to 26 °C with abundant rain throughout the year (García 1998). The site was traditionally managed, and referred to as “milpa” by the indigenous inhabitants. It is a diverse agroecosystem (polycultures) where the main crop is maize. Milpas normally contain native maize types (criollos), always accompanied by other crop plants such as bean (Phaseolus spp.) and pumpkin (Cucurbita spp.); these products constitute the traditional pre-Hispanic agriculture “food triad” system. Other common crops associated with this polyculture system are jicama (Pachyrhizus erosus) and cassava (Manihot esculenta) (López-Cano & Castillo-Campos 2009, Negrete-Yankelevich et al. 2013). Soil cores were taken at the end of the rainy season during December 2003 and January 2004. The soil is an umbric Andisol, composed of 66 % sand, 11.5 % clay, and 22.5 % lime. Other soil characteristics include 11.51 % organic matter, 9–19 mg g−1 N, 0.44 mg g−1 P, 0.21 mg g−1 K, 4.62 mg g−1 Ca, 2.00 mg g−1 Mg, and pH 5.3 (García et al. 2009).
The composite soil sample (LM38-BGBD), was a mixture of 12 subsamples taken with a corer of 5 cm diam × 20 cm height around 2 concentric circles of 3 and 6 m diam, which gave a total sampling area of 114 m2. The soil of the 12 subsamples was crumbled by hand and mixed. The sample was kept in plastic bags and stored at 4 °C, before spore isolation. The AMF spores present in the sample were morphologically identified according to INVAM keys and the manual of Schenck & Pérez (1990). The same soil samples were also used to inoculate “AMF trap cultures” with Brachiaria comata (Poaceae). This soil sample (LM28) formed part of the Conservation and Sustainable Management of Below Ground Biodiversity project (CSM-BGBD) of the Los Tuxtlas Biosphere Reserve (https://doi.org/www3.inecol.edu.mx/csmbgbd/index.php/el-proyecto-bgbd-en-mexico), and is also part of a larger sampling programme of the Lopez-Mateos community at Los Tuxtlas (Varela et al. 2009).
AMF pot cultures
The same soil samples were used to inoculate “AMF trap cultures” with Brachiaria comata (Graminae/Poaceae) using the techniques of Sieverding (1991). This plant was chosen due to its wide range of interaction with AMF, as previously reported (de Souza et al. 2004). Greenhouse conditions in Xalapa, Veracruz, were set to be similar to those at the sampling site (25–35 °C and 60–80 % relative humidity, with natural daylight and photoperiod). Pots contained autoclaved sterile sand (15 lb, 121 °C, 1 h) as a substrate, and were either irrigated with distilled water every three days, or whenever necessary to preclude drying out. A modified Hoagland’s solution (Hoagland & Arnon 1950), containing 20 µM phosphate, was used to fertilise trap cultures once or twice per month, according to the plant developmental stages. Irrigation was stopped after full development of the grass (approx. 3–4 mo), to allow for plant drying and spore production. The Brachiaria comata plants were then removed from the pots and a fresh culture was started in a new pot after approx. 3 mo (Burrows & Pfleger 2002). We considered each subculture to be one cycle (Fig. 1). Re-inoculation of pots containing 1 kg of sterile sand was carried out by using ∼40gof mixed roots with spores and sand.
Morphological identification of AMF from recently sampled field soils was performed in March of 2004 (Varela et al. 2009). Trap culture sampling of roots and spores and morphological identification of AMF took place between 2004 and 2012, for a total of nine morphological analyses. Morphological characteristics of sporocarps, spores and root subcellular structures were first observed in water on a glass slide under a dissecting microscope; specimens were later mounted in either polyvinyl alcohol:lactic acid:glycerol (PVLG; 1:1:1; Koske & Tessier 1983), or in a mixture 1:1 of PVLG and Melzer’s reagent (lactic acid/water 1:1; Brundett et al. 1994). The morphology of the spores was analyzed according to the INVAM (International Culture Collection of Vesicular-Arbuscular Mycorrhizal Fungi, https://doi.org/www.invam.caf.wvu.edu) data, and following Schenck & Pérez (1990). Photographs were taken with a digital camera (Motic digital microscope DMB3-223) on a dissecting microscope (Nikon SMZ-2T). Air-dried sporocarps and specimens mounted in either PVLG or a mixture of PVLG and Melzer’s reagent were deposited in XAL (Xalapa, Instituto de Ecología A. C., INECOL).
DNA extraction, PCR amplification and sequencing
In order to confirm the final composition of morphotypes (trap culture cycles 13 through 15) we conducted rDNA surveys (∼ 20 clones each). AMF colonized roots were cut into 0.5 cm pieces and collected in a 1.5 mL Eppendorf tube. Roots were homogenised in DNAzol® using a pestle. Genomic DNA was extracted in DNAzol® following the manufacturer’s instructions. The concentration of genomic DNA was estimated by fluorometry using the Quantit DNA Quantitation kit™ (Cat. No. Q32854, ORE).
Genomic DNA was used for molecular identification of AMF in roots, by a nested PCR protocol. The oligonucleotides used are described by Redecker et al. (2000a, b). The mixture for the first PCR reaction contained diluted DNA (1:100 v/v), 1× reaction buffer, 1.5 mM MgCl2, 0.5 µM of each primer (NS5/ITS4), 0.2 mM of each dNTP, and 1.0 unit of recombinant Taq DNA polymerase (Cat. No. 11615-010, Invitrogen, Mexico City) in 25 µL total volume. The DNA templates were initially denatured at 95 °C for 4 min. The subsequent cycles included a 60 s denaturation step at 94 °C, 60 s of primer annealing at 55 °C, and a 3 min extension step at 72 °C. A final 5 min extension step at 72 °C concluded the programme after 32 cycles. PCR was performed using a MultiGene Thermal Cycler (Model TCP600-G, Labnet International). The first PCR product was diluted 1:10 v/v with ultrapure distilled water. The dilutions were then used as DNA template for a nested PCR reaction, with primer pairs that amplify different ribosomal DNA regions (LETC1670/ITS4; ITS-1F/GIGA 5.8). The same PCR conditions were used as in the first reaction. Nested PCR bands that were ∼ 650 bp and ∼ 330 bp were cut out from a 1 % agarose gel, and the DNA extracted with the QIAquick gel extraction kit (Qiagen, Cat. 28704). The purified PCR products were cloned into the pGEM-T Easy Vector II System (Promega, Cat No. A3600, Madison, WI) as described by the manufacturer. Twenty clones were selected to make plasmid minipreps, according to the manufacturer’s instructions (QIAprep spin miniprep kit, Qiagen Cat. No. 27106, Mexico City). The amplified and cloned fragments were sequenced using the Taq Dye Deoxy Terminator Cycle Sequencing Kit (Applied Biosystems, Foster City, CA). PCR products were sequenced in both directions with an ABI Prism 3100 automated DNA sequencer (Applied Biosystems, Mexico City). Sequencing was performed using the T7 and SP6 primers. DNA sequences obtained were cleansed of
vector sequences, and screened for chimeras, which were excluded from further analysis. The sequences were queried in a BLASTN search in the NCBI database (https://doi.org/www.ncbi.nlm.nih.gov/; BLASTN v. 2.2.22; Zhang et al. 2000). AMF nucleotide sequences derived from cycle 13 were deposited in GenBank (accession numbers: KC841638-KC841656) including one sequence (KC841638) confirming the host plant identity.
Morphological analysis of the initial native mixed soil sample (LM 38-BGBD) revealed 17 morphospecies; however, in the López-Mateos area at Los Tuxtlas, 24 morphospecies have been recognized (Varela et al. 2009). The AMF morphological analysis in the trap cultures revealed both a decrease in species spore presence and a large variability during subculturing, as compared to the native soil sample. None of the field spore species reported in the first morphological soil analysis by Varela et al. (2009) were observed at the fifth cycle of successive trap culturing. In fact, four different species were found, including Claroideoglomus etunicatum. Morphological screening at the sixth successive trap culture identified Funneliformis geosporus (reported at the initial field site soil), along with C. etunicatum and two other species not found in the field (Fig. 1).
Species from Glomerales (Fig. 2A–E) were consistently found throughout successive cycles of trap culturing. On the other hand, most species of Diversisporales that were found in the field were not found in the pot cultures, except for A. morrowiae and one A. scrobiculata morphotype (Figs. 1, 2F, G). After approximately seven years of maintaining the trap cultures (cycles 13–15), C. etunicatum emerged as the dominant species (Fig. 1). Claroideoglomus etunicatum spores displayed the characteristic globose to subglobose morphology (Fig. 2A); the spores were mostly orange-brown (0-10-90-5 according to INVAM’s colour table), and 80–188 µm diam. Subtending hyphae ranged from cylindrical to slightly flared (19.4–30.2 µm wide). Composite wall thickness (L1, L2 and L3 layers) was < 1 µm, with the innermost layer of the subtending hyphae forming a structure resembling a septum (Fig. 2A). Typical internal structures formed by C. etunicatum developed in B. comata roots during cycle 13 of our subculturing (Fig. 2H, I).
All sequences obtained from cycles 13–15 were closely related to Claroideoglomus etunicatum and showed high percentages of identity (97–99.8 %) with previously reported sequences from conspecific specimens deposited in the BEG. Sequences obtained from cycle 13 were deposited in GenBank (GenBank Accession nos. KC841639–KC841656).
In this study, no Claroideoglomus etunicatum spores were found in the native field soil, although they became abundant in the pot cultures. A similar phenomenon has been reported at several sites (INVAM website: https://doi.org/invam.caf.wvu.edu/), in which AMF are not expressed in trap cultures at the beginning of subculturing, and sometimes require up to two or three subcultures to sporulate. Claroideoglomus etunicatum sporulation was inconsistent and the spores present in the soils were not found during several intermediate cycles (e.g. 7–11), although cycles 13–15 were defined by sporulation from this single species. Claroideoglomus etunicatum can be considered highly competitive, due to its ability to adapt to extremely different conditions compared to the original sampling sites. The original soil contained 11.51 % organic matter, and by using sterile sand in the trap cultures it changed to an inert substrate having no organic matter and the pH dropped from 5.3 to 4.5. Claroideoglomus etunicatum is reported to be tolerant to host plant change, which is an important factor since hosts direct the carbon input to the fungus (Manoharan et al. 2008). This species was maintained even after prolonged subculturing in the host plant, preserving its infectivity and sporulation capabilities. These observations are consistent with C. etunicatum belonging to a select group of AMF species able to survive in a single host plant (Johnson et al. 2005). This species is also reported as one of the most common and ubiquitous AMF species worldwide (Bentivenga & Hetrick 1992).
Our molecular studies confirmed the dramatic drop (by cycle 13) in species richness reported by the morphological observations of the initial soil sample. Others have previously reported that field spores found in their native soil cannot be maintained in trap cultures. For instance, Burrows & Pfleger (2002) demonstrated that after five cycles of trap subculturing, none of the species present in the field soil were found in the trap cultures. A similar phenomenon was reported by Stutz & Morton (1996): in comparison to the first trap subculture, only 75 % of the original species were found after three trap culture cycles. This suggests that a high proportion of AMF present in arid habitats may not sporulate in the field.
Species of Diversisporales have been reported to be more susceptible to environmental change than ones of Glomerales (Lovera & Cuenca 2007). The latter exhibit shorter sporulation times than Diversisporales, considered due to the small size of their spores (Bever et al. 1996) which allows them to exhibit enhanced sporulation (Hepper 1984). Glomerales species are considered generalists and have been found to be associated with grasslands (Oehl et al. 2003). Generally, species belonging to this order are competitive colonisers, which increases their probability of representation in most plant communities (Cordoba et al. 2001). Kennedy et al. (2002) previously confirmed the dominance of Glomerales in trap cultures.
Our study contributes to the growing body of evidence suggesting that the use of a single plant species in trap cultures encourages a decrease in the AMF diversity maintained. This observation may be extrapolated to field conditions where monoculture is practiced for many consecutive years, which could similarly have a negative effect on AMF diversity (Bainard et al. 2012, Li et al. 2010, An et al. 1993, Hijri et al. 2006).
Future work will be aimed at testing the efficiency of C. etunicatum from Los Tuxtlas on both the development of native plants and in agroecosystem productivity.
An ZQ, Hendrix JW, Hershman DE, Ferriss RS, Henson GT (1993) The influence of crop-rotation and soil fumigation on a mycorrhizal fungal community associated with soybean. Mycorrhiza 3: 171–182.
Antunes PM, Lehmann A, Hart MM, Baumecker M, Rillig MC (2012) Long-term effects of soil nutrient deficiency on arbuscular mycorrhizal communities. Functional Ecology 26: 532–540.
Bainard LD, Koch AM, Gordon AM, Klironomos JN (2012) Temporal and compositional differences of arbuscular mycorrhizal fungal communities in conventional monocropping and tree-based intercropping systems. Soil Biology and Biochemistry 45: 172–180.
Barea JM, Azcón R, Azcón-Aguilar C (2002) Mycorrhizosphere interactions to improve plant fitness and soil quality. Antonie van Leeuwenhoek 81: 343–351.
Bentivenga SP, Hetrick BAD (1992) The effect of prairie management practices on mycorrhizal symbiosis. Mycologia 84: 522–527.
Bevaer JD (2002) Host-specificity of AM fungal population growth rates can generate feedback on plant growth. Plant and Soil 244: 281–290.
Bever JD, Morton JB, Antonovics J, Schultz PA (1996) Host-dependent sporulation and species diversity of arbuscular mycorrhizal fungi in a mown grassland. Journal of Ecology 84: 71–82.
Brundett M, Melville L, Peterson L (1994) Practical Methods in Mycorrhiza Research. Waterloo, ONT: Mycologue Publications.
Burrows RL, Pfleger FL (2002) Arbuscular mycorrhizal fungi respond to increasing plant diversity. Canadian Journal of Botany 80: 120–130.
Chanway CP, Turkington R, Holl FB (1991) Ecological implication of specificity between plants and rhizosphere microorganism. Advances in Ecological Research 21: 121–169.
Clapp JP, Young JPW, Merryweather JW, Fitter AH (1995) Diversity of fungal symbionts in arbuscular mycorrhizas from a natural community. New Phytologist 130: 259–265.
Cordoba AS, Mendonça MM, Stürmer SL, Rygiewicz PT (2001) Diversity of arbuscular mycorrhizal fungi along a sand dune stabilization gradient: a case study at Praia da Joaquina, Ilha de Santa Catarina, South Brazil. Mycoscience 42: 379–387.
Cuenca G, de Andrade Z, Lovera M, Fajardo L, Meneses E, Márquez M, Machuca R (2003) Pre-selección de plantas nativas y producción de inóculos de hongos micorrízicos arbusculares (HMA) de relevancia en la rehabilitación de áreas degradadas de La Gran Sabana, estado Bolívar, Venezuela. Ecotrópicos 16: 27–40.
de Souza FA, Kowalchuk GA, Leeflang P, van Veen JA, Smit E (2004) PCR-denaturing gradient gel electrophoresis profiling of inter- and intraspecies 18S rRNA gene sequence heterogeneity is an accurate and sensitive method to assess species diversity of arbuscular mycorrhizal fungi of the genus Gigaspora. Applied and Environmental Microbiology 70: 1413–1424.
García E (1998) Modificaciones al Sistema de Clasificación Climática de Köppen: (para adaptarlo a las condiciones de la República Mexicana). Mexico City, Mexico: Instituto de Geografía, Universidad Nacional Autónoma de México
García JA, Fuentes T, Sosa V, Meza E, Negrete-Yankelevich S, Barois I, Bennack D, Okoth P (2009) Benchmark site description of the Los Tuxtlas, Veracruz, Mexico In: Below-Ground Biodiversity in Sierra de Santa Marta, Los Tuxtlas, Veracruz, Mexico (I Barois, J Huising, P Okoth, D Trejo, M de Los Santos, eds): 19–37. Xalapa, Mexico: Instituto de Ecología A.C. (INECOL).
Harley JL, Smith SE (1983) Mycorrhizal Symbiosis. New York: Academic Press.
Hepper CM (1984) Isolation and culture of VA mycorrhizal (VAM) fungi. In: VA Mycorrhiza (CL Powell & DJ Bagyaraj, eds): 95–112. Boca Raton, FL: CRC Press.
Hijri I, Sykorova Z, Oehl F, Ineichen K, Mäder P, Wiemken A, Redecker D (2006) Communities of arbuscular mycorrhizal fungi in arable soils are not necessarily low in diversity. Molecular Ecology 15: 2277–2289.
Hoagland DR, Arnon DI (1950) The water culture method for growing plants without soil. [Circular no. 347.] Berkeley, CA: California Agricultural Experiment Station.
Johnson D, Ijdo M, Genney RD, Anderson CI, JI (2005) How do plants regulate the function, community structure, and diversity of mycorrhizal fungi? Journal of Experimental Botany 56: 1751–1760.
Kennedy LJ, Tiller RL, Stutz JC (2002) Associations between arbuscular mycorrhizal fungi and Sporobolus wrightii in riparian habitats in arid southwestern North America. Journal of Arid Environments 50: 459–475.
Koske RE, Tessier B (1983) A convenient, permanent slide mounting medium. Newsletter of the Mycological Society of America 34: 59.
Law R, Lewis DH (1983) Biotic environments and the maintenance of sex-some evidence from mutualistic symbiosis. Biological Journal of the Linnean Society 20: 249–276.
Leal PL, Stürmer SL, Siqueira JO (2009) Occurrence and diversity of arbuscular mycorrhizal fungi in trap cultures from soils under different land use systems in the Amazon, Brazil. Brazilian Journal of Microbiology 40: 111–121.
Li LF, Li T, Zhang Y Zhao, ZW (2010) Molecular diversity of arbuscular mycorrhizal fungi and their distribution patterns related to host-plant sand habitats in a hot and arid ecosystem, southwest China. FEMS Microbiology Ecology 71: 418–427.
López-Cano EB, Castillo-Campos G (2009) Report of the vascularflora characterization and α- and β- diversity index in three communities of Los Tuxtlas, Veracruz, México. In: Below-Ground Biodiversity in Sierra de Santa Marta, Los Tuxtlas, Veracruz, México (I Barois, EJ Huising, P Okoth, D Trejo, M de Los Santos, eds): 41–47. Xalapa, Mexico: Instituto de Ecología A.C. (INECOL).
Lovera M, Cuenca G (2007) Diversidad de los hongos micorrízico arbusculares (HMA) y potencial micorrícico del suelo de una sabana natural y una sabana perturbada, Venezuela. Interciencia 32: 108–114.
Manoharan PT, Pandi M, Shanmugaiah V, Gomathinayagam S, Balasubramanian N (2008) Effect of vesicular arbuscular mycorrhizal fungus on the physiological and biochemical changes of five different tree seedlings grown under nursery conditions. African Journal of Biotechnology 7: 3431–3436.
Merckx VSFT, Janssens SB, Hynson NA, Specht CD, Bruns TD, Smets EF (2012) Mycoheterotrophic interactions are not limited to a narrow phylogenetic range of arbuscular mycorrhizal fungi. Molecular Ecology 21: 1524–1532.
Negrete-Yankelevich S, Maldonado-Mendoza IE, Lázaro-Castellanos JO, Sangabriel-Conde W, Martínez-Álvarez JC (2013) Arbuscular mycorrhizal root colonization and soil P availability are positively related to agrodiversity in Mexican maize polycultures. Biology and Fertility of Soils 49: 201–212.
Oehl F, Sieverding E, Ineichen K, Mäder P, Boller T, Wiemken A (2003) Impact of land use intensity on the species diversity of arbuscular mycorrhizal fungi in agroecosystems of Central Europe. Applied and Environmental Microbiology 69: 2816–2824.
Oehl F, Sieverding E, Ineichen K, Mäder P, Dubois D, Boller T, Wiemken A (2004) Impact of long-term conventional and organic farming on the diversity of arbuscular mycorrhizal fungi. Oecologia 138: 574–583.
Oehl F, Sieverding E, Ineichen K, Ris EA, Boller T, Wiemken A (2005) Community structure of arbuscular mycorrhizal fungi at different soil depths in extensively and intensively managed agroecosystems. New Phytologist 165: 273–283.
Oliveira RS, Boyer LR, Carvalho MF, Jeffries P, Vosatka M, Castro PML, Dodd JC (2010) Genetic, phenotypic and functional variation within a Glomus geosporum isolate cultivated with or without the stress of a highly alkaline anthropogenic sediment. Applied Soil Ecology 45: 39–48.
Redecker D, Morton JB, Bruns TD (2000a) Ancestral lineages of arbuscular mycorrhizal fungi (Glomales). Molecular Phylogenetics and Evolution 14: 276–284.
Redecker D, Morton JB, Bruns TD (2000b) Molecular phylogeny of the arbuscular mycorrhizal fungi Glomus sinuosum and Sclerocystis coremioides. Mycologia 92: 282–285.
Sanders IR, Fitter AH (1992) Evidence for differential responses between host-fungus combinations of vesicular-arbuscular mycorrhizas from a grassland. Mycological Research 96: 415–419.
Schenck NC, Perez Y (1990) Manual for the Identification of Mycorrhizal Fungi. 3rd edn. Gainesville, FL: Synergistic Publications.
Sieverding E (1991) Vesicular-arbuscular Mycorrhiza Management in Tropical Agrosystems. Eschborn: Deutsche Gesellschaft fur Technische Zusammenarbeit (GTZ).
Sieverding E, Oehl F (2005) Are arbuscular mycorrhizal fungal species invasive-derived from our knowledge about their distribution in different ecosystems? BCPC Symposium Proceedings 81: 197–202.
Smith SE, Read DJ (1997) Mycorrhizal Symbiosis. 2nd edn. London: Academic Press.
Stutz JC, Morton JB (1996) Successive pot cultures reveal high species richness of arbuscular endomycorrhizal fungi in arid ecosystems. Canadian Journal of Botany 74: 1883–1889.
Varela L, Trejo D, Álvarez-Sánchez FJ, Barois I, Amora-Lazcano E, Guadarrama P, Lara L, Olivera D, Sánchez-Gallén I, Sangabriel W, Zulueta R (2009) Land use and diversity of arbuscular mycorrhizal fungi in Mexican tropical ecosystems: Preliminary results. In: Below-Ground Biodiversity in Sierra de Santa Marta, Los Tuxtlas, Veracruz, Mexico (I Barois, J Huising, P Okoth, D Trejo, M de Los Santos, eds): 99–112. Xalapa, Mexico: Instituto de Ecología A.C. (INECOL).
Yang H, Zang Y, Yuan Y Tang J, Chen X (2012) Selectivity by host plants affects the distribution of arbuscular mycorrhizal fungi: evidence from ITS rDNA sequence metadata. BMC Evolutionary Biology 12: 50.
Yao Q, Gao J-L, Zhu H-H, Long L-K, Xing Q-X, Chen J-Z (2010) Evaluation of the potential of trap plants to detect arbuscular mycorrhizal fungi using polymerase chain reaction-denaturing gradient gel electrophoresis analysis. Soil Science and Plant Nutrition 56: 205–211.
Zhang Z, Schwartz S, Wagner L, Miller W (2000) A greedy algorithm for aligning DNA sequences. Journal of Computational Biology 7: 203–214.
We are grateful to Lucia Varela-Fregoso and Fritz Oehl for generously sharing their knowledge of AMF taxonomy, to Jesús Damián Cordero-Ramí rez for help with sequence submission to GenBank, and Jacob I. Bañuelos-Trejo for assistance with the figures. This research was partially funded by the FOMIX 94427 project (CONACYT-Veracruz Government) as part of the BioPop project; and by the agencies GEF (Global Environment Facility), TSBF (Tropical Soil Biology and Fertility), CIAT (International Center for Tropical Agriculture), and UNEP (United Nations Environment Programme), supporting initial sampling and inventories as part of the project Conservation and Sustainable Management of Below Ground Biodiversity (CSM-BGBD). We also acknowledge ECOS-France/ANUIES-CONACYT project (M08A01) for travelling and training support of Wendy Sangabriel-Conde. We thank Brandon Loveall (Improvence) for English proofreading of the manuscript.
About this article
Cite this article
Trejo-Aguilar, D., Lara-Capistrán, L., Maldonado-Mendoza, I.E. et al. Loss of arbuscular mycorrhizal fungal diversity in trap cultures during long-term subculturing. IMA Fungus 4, 161–167 (2013). https://doi.org/10.5598/imafungus.2013.04.02.01
- AM fungi