Urease producing microorganisms under dairy pasture management in soils across New Zealand

Geoderma Regional 11 (2017) 78–85

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Urease producing microorganisms under dairy pasture management in soils across New Zealand

MARK

Hossein Alizadeha,⁎, Diwakar R.W. Kandulaa, John G. Hamptona, Alison Stewartb, David W.M. Leungc, Yasmine Edwardsd, Carol Smithe a

Bio-Protection Research Centre, Lincoln University, Lincoln 7647, New Zealand Scion, Rotorua 3046, New Zealand c School of Biological Sciences, University of Canterbury, Christchurch 8140, New Zealand d School of Biological Sciences, University of Auckland, Auckland 1142, New Zealand e Faculty of Agriculture and Life Sciences, Lincoln University, Lincoln 7647, New Zealand b

A R T I C L E I N F O

A B S T R A C T

Keywords: Urease producing microorganisms Ryegrass Urea fertiliser Nitrogen Inceptisol Entisol

Urea, the most commonly used nitrogen fertiliser in New Zealand, can be quickly lost from the system via ammonia volatilisation or nitrate leaching following hydrolysis of urea by urease producing soil microorganisms (UPSMs). This study investigated UPSMs involved in urea degradation for upcoming research to reduce soil urease activity. Soils from under dairy pasture management across New Zealand, with a pasture species component of ryegrass (Lolium perenne L.) and white clover (Trifolium repens L.) and aged between 9 months to 60 years old, were collected, and UPSMs were isolated and identified using both polymerase chain reaction (PCR)-based molecular and conventional methods. The fungal genera belonged to diverse taxonomical groups including the phylum Ascomycota: class: Dothideomycetes, Eurotiomycetes, Leotiomycetes and Sordariomycetes, the phylum Basidiomycota: class: Tremellomycetes and the phylum Zygomycota: order: Mucorales, all of which have a role in urea degradation in soil. Pasture soil-resident urease producing bacteria belonged to the Gammaproteobacteria and Betaproteobacteria. Cupriavidus sp. and Mucor hiemalis showed strong urease activity when cultured on urease medium. This is the first report on the urease activity of the pasture soil inhabitants Pochonia bulbillosa, Mariannaea elegans and Gliomastix sp. This study was part of a larger study underway to investigate control of UPSMs in soil to improve the efficiency of urea utilisation.

1. Introduction Nitrogen (N), a key nutrient for the grass component of dairy pastures, is commonly applied as urea. However, urea N can be quickly lost from the system via ammonia volatilization or nitrate leaching because of urea hydrolysis following the activities of urease producing soil microorganisms (Juan et al., 2009). N availability to the plant is therefore reduced, and the subsequent production of nitrous oxide and leaching of soil nitrate are contributors to environmental damage (Denier van der Gon and Bleeker, 2005). A correlation between lowered nitrous oxide emissions and a low concentration of soil ammonium nitrogen (which is a product of urea hydrolysis by urease) in wheat fields has been demonstrated (Jiang et al., 2015). Urease (urea amidohydrolase; EC 3.5.1.5) is a nickel-dependent enzyme which can catalyse the conversion of urea to ammonia and carbon dioxide (Sirko and Brodzik, 2000), although the reason why the enzyme requires nickel but not other metals is not understood (Carter ⁎

et al., 2009). Urease was the first enzyme that was crystallised (in 1926) from jack bean and its substrate, urea, was also the first organic molecule ever synthesised in a laboratory (in 1828) (Sirko and Brodzik, 2000). Urease can be found in plants, fungi, bacteria and some invertebrates, but not in animals (Carlini and Polacco, 2008; Polacco and Holland, 1993; Sirko and Brodzik, 2000; Smith et al., 1993). In general, urease enables the organism to use urea as a nitrogen source. In some bacteria, such as Klebsiella aerogenes, urea hydrolysis has enabled them to use urea as a sole source of nitrogen (Sirko and Brodzik, 2000). Urea is also assimilated by urease in plants (Sirko and Brodzik, 2000). In many human pathogens (e.g. Proteus mirabilis, Yersinia enterocolitica, Staphylococcus saprophiticus, Helicobacter pylori and Ureaplasma urealiticum), urease can act as a virulence factor, and this has been used as a simple test to identify virulent isolates. Hydrolysis of urea by Helicobacter pylori, the agent of peptic ulceration, in the stomach, is a method for this bacterium to increase pH of the acidic environment and colonise it (Carter et al., 2009; Follmer et al., 2004; Mobley et al., 1995;

Corresponding author. E-mail address: [email protected] (H. Alizadeh).

http://dx.doi.org/10.1016/j.geodrs.2017.10.003 Received 14 July 2017; Received in revised form 10 October 2017; Accepted 12 October 2017 2352-0094/ © 2017 Elsevier B.V. All rights reserved.

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Sirko and Brodzik, 2000). Recently, it was demonstrated that blocking urease could improve plant tolerance to salt during seed germination as a result of reduced ammonium production (Bu et al., 2015). Soil urease activity can be affected by factors including moisture, temperature, soil organic matter, pH and the rate of fertiliser applied (Carter et al., 2009). A number of microorganisms have been reported to produce urease (Table 1). Around 90% of applied urea can be hydrolysed in the soil within 1–2 days of the application (Hojito et al., 2010). The half-life of urea degrading spontaneously (even at 57 °C) is around 3.6 years, (Hasan, 2000) but in the presence of urease it can be degraded 10,000 times faster (Amtul et al., 2002) suggesting a need for urease inhibitors. A number of enzymatic chemical inhibitors have been developed (Mobley et al., 1995). The use of Agrotain, a commercial urease inhibitor, in New Zealand ryegrass pastures has been reported to increase N uptake and herbage dry matter (Dawar et al., 2011). New Zealand's pastoral production industries are heavy users of nitrogen (N) fertilisers (Moir et al., 2007). Despite the importance of urea and the environmental implications of its degradation in agricultural soils, the potential contribution of the urease producing microorganisms inhabiting pasture soils has not been investigated. The current study was designed to isolate and identify New Zealand's urease producing soil microorganisms. The results can be used for future studies to work on urease reduction strategies in pastures.

Table 1 Urease producing microorganisms. No.

Scientific name

A. Fungi, actinomycetes and yeast species 1 Absidia sp. 2 Alternaria tenuissima 3 Aspergillus flavus, A. fumigatus, A. niger 4 Aureobasidium pullulans 5 Botrytis cinerea 6 Cladosporium herbarum 7 Coccidioides immitis 8 Cochliobolus heterostrophus 9 Coprinus sp. 10 Cryptococcus neoformans 11 Emericella nidulans 12 Fusarium nivale 13 Geomyces destructans 14 Humicola grisea 15 Magnaporthe oryzae 16 Malassezia furfur 17 Mucor racemosus 18 Mycosphaerella graminicola 19 Nectria haematococca 20 Neurospora sp., N. crassa 21 22 23 24 25 26 27 28 29 30 31

Paecilomyces silvatica Penicillium spp., P. brevicompactum, P. notatum Rhizopus oryzae Saccharomyces cerevisiae Saccharomycopsis Malanga Schizosaccharomyces pombe Sepedonium chrysospermum Stagonospora nodorum Streptomyces aureofaciens Trichophyton mentagrophytes Ustilago spp.

B. Bacteria 32 Anabaena cycadeae 33 Arthrobacter crystallopoietes 34 Bacillus lentus, B. subtilis, B. sphaericus 35 Bordetella pertussis 36 Brevibacterium ammoniagenes, B. stationis 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51

Burkholderia sp. Corynebacterium glutamicum Enterobacter cloacae Helicobacter pylori Klebsiella aerogenes Kocuria marina Lactobacillus reuteri Lysinibacillus sphaericus Morganella morganii Mycobacterium tuberculosis Nitrosomonas sp. Nitrosospira sp. Proteus mirabilis Providencia vermicola Pseudomonas aeruginosa, P. fluorescens

52 53 54 55

Rhodobacter capsulatus Sarcina urea Sinorhizobium medicae Sporosarcina pasteurii, S. ureae

56 57 58 59

Staphylococcus saprophiticus Streptococcus salivarius Thiocapsa roseopersicina Ureaplasma urealyticum (with the highest urease activity demonstrated) Vibrio parahaemolyticus Yersinia enterocolitica, Yersinia pestis

60 61

Reference

Donnison et al., 2000 Hasan, 2000 Hasan, 2000 & Smith et al., 1993 Hasan, 2000 Strope et al., 2011 Hasan, 2000 Sirko and Brodzik, 2000 Strope et al., 2011 Hasan, 2000 Hasan, 2000 Hasan, 2000 Hasan, 2000 Reynolds and Barton, 2014 Hasan, 2000 Strope et al., 2011 Hasan, 2000 Hasan, 2000 Strope et al., 2011 Strope et al., 2011 Hasan, 2000 & Strope et al., 2011 Hasan, 2000 Hasan, 2000 Strope et al., 2011 Hasan, 2000 Hasan, 2000 Sirko and Brodzik, 2000 Hasan, 2000 Strope et al., 2011 Hasan, 2000 Hasan, 2000 Hasan, 2000

2. Materials and methods 2.1. Soil sample collection and processing Soil samples were obtained from ryegrass (Lolium perenne L.) - white clover (Trifolium repens L.) based pastures under dairy management (hereafter referred to as pasture soils) in New Zealand located in Auckland, Canterbury, Manawatu, Marlborough, Nelson, Otago, Taranaki, Waikato, Wairarapa and West Coast (Table 2). To take samples, a straight sampling line (mostly a diagonal from corner to corner of a field) was selected and along the line six samples of similar size (400 g) were taken using a soil corer to a depth of 15 cm. The soil samples were thoroughly mixed and approximately 1 kg soil for each sample at each sampling time was placed in a sealed bag which was transferred to the laboratory and kept in a refrigerator until used for the experiments.

Hasan, 2000 Wen et al., 2015 Hasan, 2000 Strope et al., 2011 Burbank et al., 2012 & Hasan, 2000 Strope et al., 2011 Wen et al., 2015 Kang et al., 2015 Carter et al., 2009 Mobley et al., 1995 Wen et al., 2015 Hasan, 2000 Burbank et al., 2012 Mobley et al., 1995 Strope et al., 2011 Hasan, 2000 Hasan, 2000 Carter et al., 2009 Burbank et al., 2012 Wen et al., 2015 & Strope et al., 2011 Hasan, 2000 Hasan, 2000 Strope et al., 2011 Burbank et al., 2012 & Mobley et al., 1995 & Wen et al., 2015 Mobley et al., 1995 Hasan, 2000 Hasan, 2000 Mobley et al., 1995

2.2. Urease detection medium To detect microbial urease activity, a urease detection medium was prepared according to Christensen (1946) and MacFaddin (2000) with some modifications. Briefly, 1 g peptone, 1.5 g dextrose, 1.2 g sodium chloride, 0.016 g phenol red, 0.1 g nickel (II) sulphate, and 0.8 g monopotassium phosphate were dissolved in distilled water, pH was adjusted to 6.8 and after adding 18 g agar to the medium, the final volume was made to 800 ml and autoclaved. Urea (20 g product) was separately dissolved in distilled water up to a volume of 200 ml. The solution was sterilised using a Millipore membrane (0.22 μm sterile filter units), added to the autoclaved medium prior to agar setting (45 °C) and dispensed to 8 cm Petri plates. 2.3. Isolation of UPSMs from soil One gram of each soil was placed in 9 ml sterile water in a Universal container and shaken for 10 min. A serial dilution was prepared and 100 μl from each dilution was spread on urease detection medium. The Petri plates were incubated at 23 °C and colonies with a pink zone (Christensen, 1946) were isolated, cultured and purified (fungi using either a single spore or hyphal tip method; Rangaswami and Bagyaraj,

Sirko and Brodzik, 2000 Strope et al., 2011 & Mobley et al., 1995

79

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Table 2 Soil samples and types collected from different locations. Sample no.

Location

Soil series and soil type

New Zealand soil classification (Hewitt, 2010)

Soil taxonomy (Soil Survey Staff, 2014)

Pasture age

1

Pukekohe - Auckland

Typic to Mottled Orthic Granular

Typic Haplohumult

3 years

2 3 4 5 6 7

Ashley Dene - Canterbury Leeston - Canterbury Lincoln - Canterbury Oxford - Canterbury Southbridge - Canterbury Tennent Drive – Manawatu Tennent Drive – Manawatu Rai valley - Marlborough Appleby - Nelson Ettrick - Otago Hawera - Taranaki Hawera - Taranaki Te Awamutu - Waikato Greytown - Wairarapa Cape Foulwind – West Coast Sergeants Hill – West Coast

Kauaeranga deep silt loam over clay to Browns deep loam over clay Lismore stony silt loam Darnley shallow silt loam Templeton silt loam Lismore stony silt loam Waimakariri silt loam Tokomaru silt loam

Pallic Orthic Brown Typic Argillic Pallic Typic Immature Pallic Typic Argillic Pallic Weathered Fluvial Recent Argillic-fragic, Perch-gley Pallic

Lithic Dystrudept Inceptic Hapludalf Typic Dystrudept Inceptic Hapludalf Typic Dystrudept Typic Fragiaqualf

1 year 5 years 6 years 8 years 4 years 3 years

Ohakea silt loam

Mottled Immature Pallic

Typic Endoaquept

3 years

Rai silt loam Waimea silt loam Barrhill moderately deep silt loam Egmont brown loam Egmont ash loam Mairoa deep clay Greytown silt loam Flipped Pakaki soil

Typic Typic Typic Typic Typic Typic Typic Typic

Typic Dystrudept Typic Dystrudept Typic Haplustept Typic Hapludand Typic Hapludand Typic Hapludand Humic Dystrocrept Aquic Udifluvent

14 years 7 years 9 months 1 year 60 years 1 year 8 years 4 years

Kumara silt loam

Silt-mantled Perch-gley Podzol

Typic Alaquod

50 years

8 9 10 11 12 13 14 15 16 17

2005, and bacteria using the four flame streaking method; Solanki et al. (2012)) and later retested for urease activity.

Orthic Brown Fluvial Recent Immature Pallic Orthic Allophanic Ortic Allophanic Ortic Allophanic Fluvial Recent Gley Raw

solution (2% w/v) and incubated for 30 min at 37 °C before the reaction was stopped by adding 500 μl Nessler's reagent. The absorbance was read at 450 nm (using a Genesys 10 UV scanning spectrophotometer Thermo Scientific, Helios gamma) after adding 500 μl distilled water and the increase in absorbance relative to the buffer control was considered as urease activity. One urease unit (UU) was expressed as the amount of enzyme that increased absorbance by 0.1 unit at the aforementioned wavelength (Yasmeen et al., 2012).

2.4. Urease production confirmation Microorganisms isolated in 2.3 were cultured on urease production medium. Ingredients included 7 g yeast extract, 5 g bacteriological peptone, 0.1 g nickel (II) sulphate, 1 g glucose, 0.1 g sodium chloride, 4.5 g monopotassium phosphate, 10.5 g dipotassium phosphate and 0.2 g magnesium sulphate. All ingredients were dissolved in distilled water and after adjusting the pH to 6.2 the final volume was made to 1 l, distributed in tissue culture jars (19 ml medium in each 200 ml jar) and autoclaved. Each medium was inoculated with inoculum consisting of hyphae excised from a 3-day-old fungal or bacteria from 1-day old bacterial culture on potato dextrose agar (PDA) or nutrient agar (NA) media and was incubated at 28 °C on an orbital shaker (140 rpm) for 1 or 4 days (for bacteria or fungi, respectively). Urea solution (20% w/v) was sterilised using a Millipore membrane (0.22 μm sterile filter units), and 1 ml added to the medium. The jars were incubated for two days afterwards and microbial crude protein extracted as follows:

2.5. Protein content The amount of soluble proteins in different crude extracts of microorganisms was quantified using the principle of quantitative binding of proteins (Bradford, 1976) with Coomassie Brilliant Blue dye. Bovine serum albumin was used as a standard. 2.6. Total genomic DNA extraction Isolated urease producing fungi and bacteria were cultured on PDA and NA media, respectively. The plates were incubated at 23 °C and young growing colonies were used for DNA extraction. A method based on the chelation of components other than nucleic acids using Chelex® was applied to extract DNA from the microorganisms (Hennequin et al., 1999; Walsh et al., 2013).

2.4.1. Protein extraction The culture was centrifuged at 10,000g and 4 °C for 10 min. The supernatant obtained was discarded and the precipitated microorganism was washed twice with 20 mM Tris-HCl buffer and 1 mM EDTA (at pH 7) before protein extraction. The microorganism biomass was mixed with glass powder and 1.8 ml of the aforementioned buffer placed in screw cap vials and ground using a FastPrep®-24 grinder (M.P. Biomedicals, Irvine, California, USA) for 30 s. The homogenate was then centrifuged at 10,000g and 4 °C for 10 min. The supernatant thus obtained can be referred to as a microorganism intracellular protein extract (Guimaraes et al., 2006). The extracts were mixed with acetone (2:1 v/v) and incubated at −20 °C for 4 h. Pellets obtained after centrifugation were air dried and resuspended in Tris-HCl buffer (2 ml) to be later checked for ureolytic activity.

2.6.1. Fungi Mycelium (5–10 mg) of each 3-day-old fungal species growing on the PDA surface was scraped using an inoculation loop after adding 500 μl Chelex® suspension (5% w/v, in ultrapure distilled water) to the colony (Sepp et al., 1994). The suspension from each colony was collected using a micropipette, transferred to an Eppendorf tube and exposed to 1 ml liquid nitrogen. The samples were manually ground using micro pestles and then heated in boiling water for 10 min. The tubes were centrifuged at 14000g (10 min) and each supernatant was transferred to a new tube, then stored at -20 °C until they were used as the DNA template (Mohlenhoff et al., 2001). The optical density of the extracts containing total genomic DNA was measured using a Nanodrop® spectrophotometer at the wavelength of 260 nm for the DNA and 280 nm for proteins. The purity of DNA was estimated by OD 260/OD 280 and the quantification expressed as ng/μl.

2.4.2. Urease assay Tris-HCl buffer 20 mM, 1 mM EDTA (pH 7) was prepared and used for this study. To check urease activity each extract was diluted with 580 μl buffer to contain 20 μg soluble proteins, mixed with 300 μl urea 80

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chopped up and boiled in 500 ml distilled water for 30 min. The product was then filtered through a Whatman filter (No 3) and 200 ml of the extract plus 5 g sucrose, 1 g yeast and 30 g agar were mixed and taken to the final volume of 1 l using distilled water. The medium was autoclaved before dispensing in 8 cm Petri plates. Inoculum consisting of hyphae excised from a 3-day-old fungal culture on PDA medium was placed at the centre of the prune extract medium in the Petri plates and incubated at 23 °C until conidia had formed. Conventional methods based on fungal morphology were used to check the results obtained from molecular-based methods of fungal identification. Fungal morphological features were observed under a light microscope and checked with published fungal descriptions (Barnett and Hunter, 1999; Dickinson, 1968; Seifert et al., 2011; Summerbell et al., 2011). Sexual and asexual features including the colony's shape and texture, colour and growth rate on PDA; the presence, shape and colour of conidia, pigmentation, conidiophores, sporangium and pycnidia based on published fungal descriptions were considered (Promputtha et al., 2005).

2.6.2. Bacteria From each bacterium, a young colony was collected using an inoculation loop and suspended in 500 μl Chelex suspension (5% w/v). The samples were heated in boiling water for 10 min, centrifuged at 14000g (10 min) and each supernatant was transferred to a new tube which was then stored at −20 °C for later use as DNA template. DNA sample concentrations were determined using a Nanodrop® spectrophotometer. 2.7. PCR (polymerase chain reaction) amplification 2.7.1. Fungi To identify the isolated fungi to the species level, the internal transcribed spacer (ITS) region of the ribosomal operon was amplified using a standard polymerase chain reaction protocol. The amplification primers used included ITS1F: 5′-TCCGTAGGTGAACCTGCGG and ITS4R: 5′-TCCTCCGCTTATTGATATGC (White et al., 1990). The PCR mixture contained 2.5 μl PCR buffer (10 ×), 2 μl dNTP mix (2.5 mM), 1 μl of each primer (one forward and one reverse 10 μM), ultrapure distilled water 16.25 μl, 1.25 U Taq DNA polymerase and 2 μl template DNA with a total volume of 25 μl for each reaction. The following temperature regime was used as the PCR programme for ITS amplification: Initial denaturation at 95 °C for 5 min, followed by 40 cycles of denaturation at 95 °C for 45 s, annealing at 57 °C for 45 s and extension at 72 °C for 2 min. The amplification was terminated by a final extension at 72 °C for 7 min after the last cycle. Five microlitre aliquots of each amplification product were separated by electrophoresis (Midicell EC350 electrophoretic gel system) on 1% agarose gels in 1 × TAE buffer (Tris-Acetate-EDTA buffer) prestained with RedSafe™ at 120 V for 45 min, visualized and photographed under UV light using a VersaDoc™ imaging system. The samples along with primers were then sent to the Lincoln University sequencing facility, Bio-Protection Research Centre (Lincoln, New Zealand) to be sequenced using an Applied Biosystems 3130xl Genetic Analyzer, with a 50 cm array and POP7 installed as the standard platform. The same primers used for sequencing and sequence files generated from ITS sequencing were edited and assembled using the ChromasPro software (http://www.technelysium.com.au/ChromasPro. html) and compared to the nucleotide database at the US National Centre for Biotechnology Information (NCBI) to find the nearest relatives (http://blast.ncbi.nlm.nih.gov/Blast.cgi?CMD=Web&PAGE_ TYPE=BlastHome).

2.9. Statistical analysis Data collected from urease test were subjected to analysis of variance (ANOVA, p ≤ 0.05) using GenStat 18 software (VSN International Ltd) followed by comparison of mean values of the treatments using Fisher's unprotected least significance difference. 3. Results 3.1. Medium, isolation and urease test Urease producers were isolated from the soil samples based on the appearance of a pink colour around their colonies on the urease medium. Urease activity varied among the isolates; some possessed weak activity while some were quite strong. There was a difference in the microbial community for different locations; in Nelson, for example, soil dominant urease producers were bacteria in comparison with Oxford where they were fungi. All of the microorganisms were confirmed as being able to produce urease in the medium and extracts obtained from Cupriavidus sp. and Paecilomyces carneus showed strong specific urease activity (Fig. 1). 3.2. Identification of the isolated microbes 3.2.1. Molecular-based identification The DNA yields from most isolates using Chelex® resin were reasonably high and a sharp band was seen after gel electrophoresis of the amplification products. When 5 μl of the PCR product was run in agarose gel, a fragment size of approximately 600 bp for fungi and 1500 bp for bacteria was obtained (Fig. 2).

2.7.2. Bacteria The bacterial 16S rRNA genes were amplified using primers f8–27: 5′- AGAGTTTGATCCTGGCTCAG and r1510: 5′-GGTTACCTTGTT ACGACTT (Lipson and Schmidt, 2004). The PCR mixture contained 2.5 μl PCR buffer (10 ×), 2 μl dNTP mix (2.5 mM), 1 μl of each primer (one forward and one reverse 10 μM), ultrapure distilled water 16.25 μl, 1.25 U Taq DNA polymerase and 2 μl template DNA with a total volume of 25 μl for each reaction. PCR amplification was performed under the following conditions: 94 °C for 3 min, 30 cycles (94 °C for 1 min, 57 °C for 1 min, 72 °C for 2 min) and 72 °C for 10 min (Nguyen et al., 2011). The DNA yielded was visualized by electrophoresis in 1% (w/v) agarose gel pre-stained with RedSafe™ and the process then followed by sequencing of the resulting product at Lincoln University sequencing facility. BLAST (basic local alignment search tool) analysis for nearest relatives was performed through EzTaxon-e identification service (http://eztaxon-e. ezbiocloud.net/) after edition of the sequence files using ChromasPro software (Chun et al., 2007; Kim et al., 2012).

Fig. 1. Urease activity in intracellular extracts of urease producing microorganisms. 1Cladosporium cladosporioides, 2- Fusarium culmorum, 3- Pochonia bulbillosa, 4- Lewia infectoria, 5- Cylindrocarpon candidulum, 6- Fusarium oxysporum, 7- Phoma sp., 8Paecilomyces carneus, 9- Cupriavidus sp. Values labelled with the same letter do not differ significantly according to Duncan's multiple range test (P < 0.05).

2.8. Morphological identification of fungi Prune extract agar medium was used to induce fungal conidiation for morphological studies. Briefly, 25 g of de-stoned dried prunes were 81

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Table 3 Identification of the isolates using ITS (urease producing fungi) and 16S rRNA (urease producing bacteria) sequencing.

Fig. 2. PCR amplification of 16S rRNA genes of bacteria and ITS regions of fungi using specific primers. The amplified fragments of 16S rRNA genes (from bacteria) and ITS regions of fungi represent 1500 and 600 bp, respectively, when analysed by electrophoresis on a 1% agarose gel.

Comparison of the gene sequences obtained from the test microorganisms with the sequences available in databases revealed their relationship with various species. Microorganisms isolated from the New Zealand pasture soils sampled are presented in Table 3. Fungal species related to a diverse taxonomical group included the phylum Ascomycota: Cladosporium cladosporioides, Lewia infectoria, Phoma sp. (calss: Dothideomycetes), Paecilomyces sp., Penicillium spinulosum (Eurotiomycetes), Geomyces sp. (Leotiomycetes), Chaetomium sp., Cordyceps clamydosporia, Fusarium sp., Gliomastix sp., Humicola grisea, Mariannaea elegans, Nectria haematococca, Pochonia bulbillosa, Thelonectria veuillotiana (Cylindrocarpon candidulum) (Sordariomycetes), the phylum Basidiomycota: Cryptococcus victoriae, Trichosporon sp. (Tremellomycetes), the phylum Zygomycota: Absidia sp., and Mucor hiemalis (order: Mucorales), all of which have a role in urea degradation in soil. All pasture soil-resident urease producing bacteria belonged to Gammaproteobacteria except Cupriavidus sp. which has been categorised in Betaproteobacteria. Cupriavidus sp. and Mucor hiemalis showed strong urease activity when cultured on the urease medium. This list includes a wide range of microorganisms including saprophytes (e.g. Gliomastix protea), a human pathogen (Trichosporon sp.), insect and nematode pathogens (Paecilomyces marquandii; Pochonia bulbillosa) and plant pathogens (e.g Fusarium solani). When Cupriavidus sp. was cultured on urea medium the colour of the medium converted to pink within 2 min suggesting the bacterium's strong urease activity. Some UPSMs were widespread while others very limited. For example, Fusarium oxysporum was isolated from various locations and soil types including Auckland (Typic Haplohumult), Canterbury (Lithic Dystrudept), Otago (Typic Haplustept), Waikato (Typic Hapludand) and Wairarapa (Humic Dystrocrept) while Geomyces was only found in Lincoln (Typic Dystrudept). 3.2.2. Morphological-based identification The results obtained from molecular-based methods of fungal identification were confirmed using fungal morphological features. Examples are provided in Fig. 3. The morphology of all isolates was in agreement with the description of the species in the literature. Simple phialides producing dark-colored conidia in slimy heads were observed in Gliomastix sp., (Fig. 3A). The presence of dictyochlamydospores as a distinctive character was used to discriminate Pochonia species from Verticillium except for P. globispora which does not produce dictyochlamydospores (Zare and Gams, 2007).

Isolate no.

Sample no.

Location

Scientific name

Similarity %

1 2 3

1 1 3

Auckland Auckland Canterbury

99 99 100

4

3

Canterbury

5 6 7 8 9 10 11 12 13

3 2 4 4 4 4 5 5 5

Canterbury Canterbury Canterbury Canterbury Canterbury Canterbury Canterbury Canterbury Canterbury

14 15 16

6 7 7

Canterbury Manawatu Manawatu

17 18 19 20 21 22 23 24 25 26 27 28 29

8 8 8 8 9 9 10 10 11 11 11 11 13

Manawatu Manawatu Manawatu Manawatu Marlborough Marlborough Nelson Nelson Otago Otago Otago Otago Taranaki

30 31 32 33 34 35 36 37 38 39 40 41 42 43 44

12 13 12 12 14 14 14 14 15 15 17 17 16 16 4

Taranaki Taranaki Taranaki Taranaki Waikato Waikato Waikato Waikato Wairarapa Wairarapa West Coast West Coast West Coast West Coast Canterbury

45 46 47 48 49 50

9 9 10 10 10 12

Marlborough Marlborough Nelson Nelson Nelson Taranaki

51 52 53 54

12 14 15 16

Taranaki Waikato Wairarapa West Coast

Fusarium oxysporum Pochonia bulbillosa Cladosporium cladosporioides Cordyceps chlamydosporia Fusarium culmorum Fusarium oxysporum Humicola grisea Geomyces sp. Cryptococcus victoriae Paecilomyces carneus Absidia sp. Lewia infectoria Thelonectria veuillotiana (Cylindrocarpon candidulum) Fusarium oxysporum Fusarium solani Gibberella zeae (Fusarium graminearum) Paecilomyces marquandii Penicillium spinulosum Pochonia bulbillosa Pochonia sp. Chaetomium sp. Fusarium solani Fusarium solani Mariannaea elegans Fusarium oxysporum Gliomastix sp. Paecilomyces carneus Phoma exigua Cladosporium cladosporioides Mucor hiemalis Phoma sp. Pochonia sp. Trichosporon sp. Fusarium oxysporum Paecilomyces lilacinus Paecilomyces marquandii Phoma paspali Fusarium oxysporum Mucor hiemalis Nectria haematococca Paecilomyces carneus Phoma sp. Pochonia sp. Pseudomonas chlororaphis Rahnella aquatilis Serratia liquefaciens Citrobacter freundii Enterobacter ludwigii Pseudomonas agarici Pseudomonas azotoformans Rahnella sp. Pseudomonas sp. Serratia proteamaculans Cupriavidus sp.

100 100 100 99 98 99 97 98 99 98

99 97 100 98 99 97 96 98 99 98 96 100 95 100 98 100 99 98 98 99 99 98 98 100 99 99 99 99 97 99 99.2 96.3 99.4 97 99.8 98 99.6 99 99 99 98

2011). As, in this research, a pH change due to urea hydrolysis was used to detect urease activity, no data could be provided regarding the presence of either of the processes in the isolated microorganisms. PCR-based molecular methods have been used for fungal identification and diagnosis (Mohlenhoff et al., 2001; Schoch et al., 2012) as well as in bacterial phylogeny and taxonomy studies (Eden et al., 1991; Janda and Abbott, 2007). The primers used in this study allowed the amplification of the target regions by PCR. The score of Gliomastix was

4. Discussion Urea can be degraded by urease or urea amidolyase to produce ammonia and carbon dioxide in a one or two-step process, respectively (Strope et al., 2011). There is evidence that a urea amidolyase gene from bacteria has been horizontally transferred to fungi (Strope et al., 82

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Fig. 3. Morphological-based identification of urease producing fungi; A. Gliomastix sp. - large spore balls, B. Absidia sp. – sporangia, C. Fusarium graminearum – macroconidia, D. Humicola grisea – aleuriospore, E. Arthrospores of a human pathogen, Trichosporon sp.

Lactobacillus reuteri, Staphylococcus spp., Bacillus spp., Corynebacterium glutamicum and Streptomyces spp. in New Zealand has been previously demonstrated according to the New Zealand Organisms Register website (http://www.nzor.org.nz/search). These bacteria might not be pasture soil inhabitants, or if present, these specific isolates lacked the ability to produce urease. Some Gram negative bacteria including Sinorhizobium medicae, Bordetella spp., Burkholderia sp., Yersinia spp., Vibrio sp., Morganella morganii and Klebsiella spp. were also not isolated from the pasture soils in this study, suggesting that they are not likely pasture soil inhabitants. Other soil urease producing bacteria including Thiocapsa sp., Ureaplasma urealiticum (Gram negative), Kocuria marina (Gram positive) and a fungus Coccidioides sp. have not been previously reported in New Zealand, explaining their absence in the list of the identified microorganisms in this project. Although a number of other fungi and actinomycetes including Aspergillus (Emericella), Aureobasidium, Coprinus, Malassezia, Neurospora, Saccharomyces, Sepedonium, Trichophyton, Ustilago, Schizosaccharomyces, Botrytis, Cochliobolus, Mycosphaerella, Magnaporthe, Rhizopus and Stagonospora are present in New Zealand (New Zealand Organisms Register website) and reported as urease producers in the literature (Hasan, 2000; Sirko and Brodzik, 2000; Strope et al., 2011), none of them was isolated in this study. This could either be due to their absence from pasture soils, or that the method used for isolation was ineffective.

low and its morphological features did not match the references (Dickinson, 1968; Summerbell et al., 2011) suggesting that it could be a new species but this remains to be confirmed. This fungus has an important role in biodeterioration of wall paintings (Kiyuna et al., 2011). Different microorganisms including filamentous fungi, yeasts, mycorrhiza, bacteria, cyanobacteria and actinomycetes have been shown to produce ureolytic enzymes (Burbank et al., 2012; Carter et al., 2009; Hasan, 2000; Kang et al., 2015; Mobley et al., 1995; Sirko and Brodzik, 2000; Smith et al., 1993; Strope et al., 2011; Wen et al., 2015). The culturable microorganisms isolated in this study belonged to various categories including Ascomycota (Taphrinomycotina, Pezizomycotina and Saccharomycotina), Basidiomycota, Zygomycota, and included plant and human pathogens and also saprophytes. The urease activity of some of the isolated microorganisms, including Pochonia bulbillosa, Mariannaea elegans and Gliomastix sp., has not been previously reported but our results regarding urease activity of Cladosporium, Fusarium, Humicola, Cryptococcus, Paecilomyces, Trichosporon, Alternaria, Penicillium, Mucor, Nectria, Enterobacter and Pseudomonas are consistent with previous studies (Hasan, 2000; Kang et al., 2015; Strope et al., 2011). No Gram positive urease producing bacteria or actinomycetes were isolated from New Zealand pasture soils in this study, although the presence of a number of Gram positive bacteria and actinomycetes including Lysinibacillus spp., Streptococcus spp., Arthrobacter spp., 83

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reduce pollution resulting from urea degradation by these microbes.

Emericella nidulans was found in sewage sludge in a previous study (Hasan, 2000) and we did not isolate it in ryegrass pastures. Cupriavidus sp. showed very high ureolytic activity and started urea hydrolysis within 2 min. Rapid hydrolysis of N-urea fertiliser can cause a concomitant rise in soil pH because of ammonium accumulation, and thus damage germinating seedlings. As urease is an enzyme which is produced by soil urease producing microorganisms (either culturable or non-culturable), reduction of the population of these microorganisms in soil should reduce N losses and allow an increase in N use efficiency. This has been neglected in previous studies, which have all focused on chemical inhibitors of the enzyme (Dawar et al., 2011). Most of the isolates found in this study were fungi. High colonisation by fungi (rather than bacteria and actinomycetes) in soils treated with urea has been previously reported (Hasan, 2000) and as New Zealand dairy farmers are heavy urea users, this could explain the higher population of fungal urease producing isolates than bacterial ones. The human pathogen, Trichosporon sp., reported previously as a urease producing fungus (Hasan, 2000), was also found to produce urease in this study. This yeast, like other dermatophytes, uses the enzyme to hydrolyse urea present in sweat and this character has been used as one of the identification criteria (Hasan, 2000). Some soil-borne microorganisms can play more than one role in the process of urea hydrolysis, nitrification and denitrification. For example, some ammonia oxidising bacteria (such as Nitrosomonas sp.) and archaea have been reported to produce urease (Hasan, 2000; Lu et al., 2012), suggesting that inhibition of growth of some soil urease producing migroorganisms could directly affect the nitrification process. In this study, no research was carried out regarding the likely expression of the genes involved in nitrification. A number of genera including Fusarium, Pochonia, Cladosporium, Phoma, Pseudomonas, Rahnella and Serratia were isolated from both the North and South Islands, while some including Cordyceps, Humicola, Geomyces, Cryptococcus and Mariannaea were only found in the South Island (refer to KMZ file). Soils were collected from pastures which ranged in age between 9 months to 60 years. Most of these pastures had a silt loam texture. Phoma was isolated from both the youngest and oldest pastures in this study, suggesting this fungus is present in both Islands and can persist in soil. Some microorganisms, for example, Fusarium, Pochonia, Cladosporium, Paecilomyces, Phoma and Pseudomonas were found in different soil types, while some were in only one soil type, e.g. Chaetomium (in a Typic Dystrudept) and Trichosporon (in a Typic Orthic Allophanic). In this study we isolated and identified only culturable microbes from the soils sampled. Although the proportion of soil bacteria that are culturable is small (1–10%, Ward et al., 1990), if urease producing microorganisms are taken as a subset of the total, this does not necessarily mean that only 1–10% of urease producing microorganisms are culturable. This information is unknown, as to the best of our knowledge, no one has worked on the proportion of culturable and unculturable urease producing microbes in soil. Also even if culturable soil urease producing microbes are 10% or less of the total, this does not necessarily mean their urease producing activity is 10% or less of urease activity in the soil. Isolation and identification of the microbes from pasture soils in this study is new, and we acknowledge that it does relate only to a subset of the soil microbial community. However, our purpose was to isolate and identify culturable soil urease producing microbes from under dairy pastures, and to confirm their ability to produce urease. We did not set out to quantify urease activity in the soil. These isolates could be used for further studies related to optimisation of urea application based on the microbial favoured environmental conditions to increase nitrogen use efficiency in plants. For example, based on the population of urease producing microbes in soil, their individual activity, favoured pH, favourable soil temperature and moisture content, and climate data, it will be possible to develop a modelling system to optimise time of urea application in order to

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