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Determination of agricultural impact on soil microbial activity using # O and respiration experiments 18
P HCl
Katherine Polain, Christopher N. Guppy, Oliver Gimli Gunning Knox, Leanne Lisle, Brian Wilson, Yui Osanai, and Nina Siebers ACS Earth Space Chem., Just Accepted Manuscript • DOI: 10.1021/ acsearthspacechem.8b00021 • Publication Date (Web): 04 May 2018 Downloaded from http://pubs.acs.org on May 5, 2018
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ACS Earth and Space Chemistry
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Determination of agricultural impact on soil microbial
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activity using δ18OP HCl and respiration experiments
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Katherine Polain1*, Christopher Guppy1, Oliver Knox1, Leanne Lisle1, Brian Wilson1,
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Yui Osanai1 and Nina Siebers2
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*Corresponding author email:
[email protected] 6
Abstract
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Improved understanding of microbial activity and associated nutrient cycling in
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agricultural systems is required to maximise production whilst maintaining and
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improving soil fertility. We compared past and current microbial activity using the
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stable δ18OP HCl pool and respiration incubations, under crop and native systems to
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a depth of 1 m. Contrary to current understanding, agricultural practices have not
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decreased microbial activity in our crop system. Differences in average δ18OP
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signatures between land use systems, indicated higher past microbial activity in the
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entire soil profile under the crop system (13.7 ‰) compared to the native system
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soil profile (11.0 ‰), whilst current microbial activity was double under the crop
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system, especially between 15 and 100 cm. Evenly distributed microbial activity in
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the top and subsoils of the crop system, as well as an increase of biomass in the
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native system subsoil, highlight the importance of investigating microbial dynamics
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beyond the top 0-30 cm of the soil profile. In these relatively dry, carbon and
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nutrient poor Australian soils, the influence of water is perhaps the key to explaining
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our results.
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Abstract art
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HCl
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Keywords
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Cotton, oxygen isotopes, respiration, subsoil, vertosol
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Introduction
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As the world’s population continues to grow, soil security is of utmost importance to
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meet the demand for food, feed and fibre.1 Cotton, as the largest supplier of natural
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fibre globally, is therefore a vital industry and is under pressure to maintain and
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improve soil resources, whilst maximising yields.1 Cotton has been a significant part
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of Australia’s agricultural landscape for over 100 years and continues to evolve with
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the modern industry producing yields that are three times the world average,
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contributing over $2 billion to the nation’s economy.2, 3
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Cotton growing regions are confined to the eastern states of Australia, with most
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crops grown in vertisol (international classification) or vertosol (Australian
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classification) soils, also referred to as ‘cracking clays’.4 In contrast to European and
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North American soils, which are relatively new and fertile, Australian soils are ancient
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and weathered. As such, the fertility of Australian soils is low by global standards,
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with low levels of biologically available carbon (C).5,
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background, the cotton growing vertisols have good fertility and water holding
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capabilities by Australian standards.4
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Vertisols in Australia have been intensively used since the latter half of the 20th
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Century for crop, particularly cotton and grains, production. Vertisols typically have
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>35% clay and exhibit a range of vertic properties including self-mulching, cracking,
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slickensides and lenticular peds.4 The high clay content of these soils influences
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their structure, drainage characteristics and therefore management. An intensive
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cropping history has made Australian vertisols inherently low in soil organic matter
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(SOM) and this limitation has been exacerbated by modern cropping practices
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resulting in lower SOM mineralisation by soil biota and subsequent reduction in C
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and secondary metabolites (such as nitrogen, N, and P) cycling.1, 7 The associated
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Against this ancient
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nutritional and management challenges that these properties pose have direct
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implications for the soil biology, with studies by Bell et al.8 finding that addition of
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inorganic fertilisers and other management practices having little or no effect on soil
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biota and subsequent nutrient cycling to make these nutrients bioavailable for crops.
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In Australian cropping systems, microbial activity and diversity is lower than any
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other land use resulting in extensive yield, soil structure and fertility deterioration,
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mostly attributed to low soil C availability.8-10 To date, much of our understanding of
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microbial activity (and subsequent links to nutrient cycling) has been undertaken in
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the top 30 cm of agricultural and native soils.8, 10-13 However, cotton roots are known
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to grow beyond this depth to at least 0.6 m, which means there is potential subsoil
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cycling and utilisation of nutrient reservoirs that has yet to be investigated to evaluate
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soil biological activity and health.14
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A powerful tool to trace microbial nutrient cycling processes, is the oxygen (O)
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isotope composition of phosphate.15 Bonding between O and P is stable under
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abiotic conditions where no exchange between oxygen isotopes and water occurs.16
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There is evidence to suggest that the natural abundance of δ18O, the ratio between
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16
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bioavailable soil phosphates (δ18OP), reflects bedrock origins where microbial and
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root enzyme activity is absent.17 In the presence of biological activity, and
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phosphate-processing enzymes there is an O exchange between water and
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phosphate. During P mineralization of organic matter (e.g., phosphomono- and
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diesterases) by extracellular enzymes, this exchange is unidirectional leading to a
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one-time shift of δ18OP.18 On the other hand, intracellular enzymes (e.g.,
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pyrophosphatase) promote repeated cycling of the phosphates and thus will lead to
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a temperature dependent equilibration between O isotopes of the ambient water and
O and
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O analysed in relation to the ratio of defined standard materials, in
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phosphate.
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turnover, the δ18OP value will deviate further from the original δ value in the
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bedrock.17 However, the resulting δ18OP value depends on several factors, e.g, the
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δ18O of the bedrock, the δ18O of the organic source, the δ18O of the ambient water,
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and the composition of the enzymes involved in P cycling.
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This principle has been used by researchers studying P cycling in soil using δ18OP
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values in bioavailable P pools of natural systems or within incubation studies by
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using
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time.20-23 Changes to δ18OP in the more stable HCl extractable-P pool are generally
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smaller in comparison to the bioavailable P pools, as HCl-P does not directly
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participate in biological P cycling.24, 25 However, conclusions on microbial P turnover
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can be drawn from the HCl-P pool, since secondary phosphate minerals precipitated
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over long time periods use phosphate that may have undergone microbial turnover in
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the soil solution, making biotic changes detectable in this pool.26, 27 During P cycling,
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some of the cycled P is precipitated into the more stable HCl-P pools and cycling
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effects are visible in HCl-P leading to a shift in δ18OP
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δ18OP measurements is ideal for Australian vertisols as the soils are old, with the
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main P form being calcium phosphates (Ca-P), which are slowly bioavailable and in
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small amounts, resulting in long term signatures that are reflective of past microbial
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activity.26, 28-30
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Present microbial activity can be assessed through the monitoring of soil respiration
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rates using an apparatus (respirometer) developed by Nordgren31. Soil respiration is
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the production of carbon dioxide (COଶ ) by soil organisms (including soil
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Hence, where there is higher biological involvement in soil P
O-labeled water to examine the enzymatic O exchange with phosphate over
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Using the HCl-P pool for
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microorganisms) and plant parts (roots and rhizomes) in the soil, where energy is
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gained through metabolising organic matter.32
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Determining long-term δ18OP
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respiration allows us to compare past and present microbial activity in both crop and
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native system, to gauge the impact of agricultural practices on Australian vertisol
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surface and subsoil health, as well as offering insight into some biological P cycling
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processes. The work presented here is part of a larger project to determine
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ecologically and economically balanced best agricultural practices, ensuring the
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Australian cotton industry continues its contribution to meeting the demands of an
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increasing global population. We hypothesised that both historic and current
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microbial activity would be higher in the native vegetation system due to less
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disturbance to the soils and their processes.
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Materials and Methods
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Site and soil sampling
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The crop site (Latitude 30o11’39.45”S, Longitude 149o36’16.2”E) was located at the
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Australian Cotton Research Institute (ACRI), 26 km north west of the Narrabri
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township, New South Wales. The native vegetation site was a further 5 km west of
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ACRI (Latitude 30o12’10.08”S, Longitude 149o32’44.88”E) and comprised of river red
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gum (Eucalyptus camaldulensis)/coolabah (Eucalyptus coolabah) open woodland
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with a redgrass (Bothriochloa macra) and prickly acacia (Vachellia nilotica)
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understorey. This region experiences a semi-arid climate with mild winters (daily
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mean 10 oC to 19 oC) and hot summers (daily mean 20 oC to 32 oC), with a mean
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average rainfall of 646 mm.33
HCl
signatures and measuring current microbial
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Both sites are alkaline dark grey cracking medium clay (vertisol, approximately 66%
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clay), classified as fine thermic montmorillonitic Typic Haplustert.
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0-1 m soil has previously been characterised as being alkaline, with an average pH
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of 7.5 (in 0.01M CaCl2) and average conductivity of 0.29 dS m-1.4,
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decreases down both soil profiles, from 1.05 to 0.60% in the crop system and 1.78 to
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1.58% in the native system. The gravimetric water content has been established as
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increasing with depth from 0.13 to 0.17 in the crop system and being 0.09 down the
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soil profile of the native system, with the exception of 0.14 in the surface soil.
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The crop site is managed since 1985 using minimum tillage, with soil disturbance
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occurring prior to planting, during pupae busting (elimination strategy used for
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Helicoverpa spp. larave) for insect management, affecting the top 10 cm of the
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profile. According to farm records, mineral P fertiliser has been applied only once to
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this long term rotational trial in the early 1990’s. The farm record did not record either
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the type or amount of P fertiliser applied, but it is believed to have been ~20 kg P ha-
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1
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the gined cotton yields over the last 10 years, the crop accumulates 35 kg P ha-1 ,
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with approximately 20 kg P ha-1 removed at harvest.34 When rainfall is insufficient,
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crop fields are irrigated at a rate of 1 ML ha-1 (equivalent to 100 mm rain) at 7-20 day
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intervals, resulting in 4-8 irrigations per season, whereas the native site is reliant on
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rainfall.
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Sampling of the crop cores took place during January 2016 (mid-season), while
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native vegetation samples were collected in May 2016. All samples were extracted
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using a powered soil corer 42 mm in diameter, collecting 4 soil cores in each system
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to a depth of 100 cm. Intact cores were recovered into polyvinyl chloride (PVC) half-
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pipes to store and support the core, sealed with plastic wrap and transported back to
4, 14, 34, 35
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The crop
Total carbon
. The native site has never been fertilised or commercially cropped. On average of
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the laboratory in cooled, insulated containers for processing. In the laboratory, soil
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cores were sub-divided into five depths (0-15, 15-30, 30-50, 50-70 and 70-100 cm).
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Irrigation water samples from January 2015 were supplied by staff from the New
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South Wales Department of Primary Industries (NSW DPI). This water was pumped
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from the river, providing δ18OW representation for both crop and native systems.
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Basalt rock samples were collected from Sawn Rocks (Latitude -30o8’42.12”,
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Longitude 150o3’12.41” and elevation 460 m), located 40 km away from the sample
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sites. These rocks were formed during the same period of volcanic activity and under
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similar conditions as the events under which the soil parent material was derived.
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Sequential extraction of P
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Soils were sequentially extracted using a modified Tiessen and Moir
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Resin and sodium bicarbonate soluble P are extremely small fractions in our soils, so
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they were omitted in favour of NaOH extraction, which incorporates these two.36
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Briefly, 0.5 g of the air-dried and sieved soil sample was mixed with 30 mL of 0.1 M
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NaOH and shaken for 16 hours on an orbital shaker at 220 rpm. Samples were
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centrifuged at 900 x g at room temperature for 30 minutes and the supernatant was
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collected for analysis. The soil pellet was resuspended in 30 mL of 1 M HCl and
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once again shaken for 16 hours and centrifuged to collect the supernatant for
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analysis. Remaining soil P was determined following the digestion method of Till,
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McArthur and Rocks37. The soil pellet from HCl extractions was digested using 10
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mL of a 1 L mixture of 0.3 % potassium dichromate, 43.4% perchloric acid, 42%
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nitric acid and 1% bromine in deionised water. The NaOH, HCl and digest extracts
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were run on a Model 725 radial viewed Inductively Coupled Plasma-Optical Emission
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Spectrometer (ICP-OES) with mass flow controller (Agilent, Australia) against known
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method.
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standards of ammonium dihydrogen phosphate (99.99%) to determine P
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concentration at a wavelength of 213.618 nm.
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Oxygen isotopic composition of phosphate (δ18OP HCl)
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Before purification and extraction, soil samples for δ18OP HCl analysis were oven-dried
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at 40 oC until a constant mass was obtained. Samples were then ground to pass a
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