Summary
We used a combination of approaches to characterize the diversity
and functions of soil microbes and different soil properties from
four microhabitats: rootzone soil under Tarbush (FLCE, a dominant
Chihuahuan Desert shrub), rootzone soil under a dominant perennial
grass (PLMU, MUPO depending on landform), biocrust, and the
uncrusted/poorly crusted soil in the plant interspace. We
collected soil samples from each of the four microhabitats at four
locations in the Jornada Experimental Range using sterile sampling
techniques. We used the phospholipid fatty acid assay (PLFA) to
assess differences in microbial biomass and microbial community
composition among the microhabitats and landforms. We used the
biologically based phosphorus (BBP), Olsen-P, and potassium
sulfate extraction methods to measure phosphorus availability in
the soils, and we applied standard methods to measure physical
soil characteristics including soil pH, texture and elemental
composition of the water-soluble fraction. Exoenzyme activity was
assessed with a modified high-throughput microplate procedure
(McLaren et al., 2017).
Topsoil sampling
At each micro habitat (Tarbush, Grass, Biocrust, Uncrusted) per
site we collected 5 soil cores at systematically placed locations
within a 25 cm2 area to a depth of 2 cm. The soil corer had a
diameter of 6 cm. Soil samples were collected during the dry
season. We used sterile sampling techniques to obtain soil
samples. The 5 core samples were combined to yield one composite
sample representing each microhabitat at each site (= 4 composite
samples per site; one for each Tarbush, Grass, Biocrust, Uncrusted
per site).
Subsurface sampling
At the same location of topsoil sampling, we also excavated a soil
pit as follows. At the vascular plant microhabitats the pit was
about 1 m x 50 cm wide for shrubs and 50 x 50 cm wide for grasses
guided by the vegetation drip line. The pit extended to a depth of
30 cm or to the top of caliche, whatever was shallower, to expose
roots with attached soil (rhizosphere). Rhizosphere and rhizoplane
(short rootzone) soil samples were collected for shrub and grass
root zones, removing ca. 50-100 g of soil and roots each soil pit.
At the biocrust and uncrusted microhabitats we excavated a 25 cm2
by 30 cm deep pit and sampled the bulk 2-20 cm soil layer as a
composite. There were 4 total excavations at each site and 12
excavations at each landform (4 micro habitats x 3 replicates x 4
landforms = 48 soil excavations). Surface and subsurface samples
were placed in a cooler, transported to the NMSU soil science
laboratory, and stored in a 4oC cool room until processing.
Standard soil physical, chemical, and biological characterization
In the laboratory, the fine earth fraction (< 2mm soil) was
obtained using a 2 mm standard soil sieve by removing litter and
gravel and gently crushing soil aggregates through the mesh. We
used sterile technique to handle and process the samples this way
(i.e., gloves and ethanol to clean metal sieve, sieve pan, and
metal handling tools). Percent gravel content was determined
gravimetrically from total sample weight. The fine earth fraction
was then well mixed and split using sterile technique in a laminar
flow hood for subsequent soil microbial, soil chemical, physical
and nutrient analyses. Gravel and litter removal as well as
subsampling was done within max 2 weeks after sampling. A 20 g
subsample was sent to Ward Laboratories, Inc. in Kearney, NE, USA
for microbial biomass and composition determination by the
Phospholipid Fatty Acid (PLFA) method (within 2 weeks after sample
collection, Findlay 2004). We determined soil pH and electric
conductivity (EC) using the saturated paste method using about
150-200 g of soil depending on texture via an Oakton Cole-Palmer
pH/CON 510 Benchtop Meter (Vernon Hills, IL, USA) (U.S. Salinity
Laboratory Staff, 1954). Soil texture was assessed using a Malvern
Mastersizer 2000 laser diffractometer (Malvern Instruments Ltd.,
Worcestershire, UK), after organic matter and particles bigger
than 2 mm were removed at Dr. Thomas Gill’s lab at UTEP. Inorganic
and organic soil carbon content were determined using a LECO SC632
Sulfur/Carbon Determinator (LECO Corporation, MI, USA).
Available nutrients in soil were extracted in 0.5 M Potassium
Sulfate (K2SO4). Soil samples (5 g fresh mass) were thawed and
extracted with 25 mL of 0.5 M K2SO4 by shaking for 2 h and then
filtered through glass filter paper. Phosphate (PO43-) in extracts
was analyzed using colorimetric microplate assays (BioTEK Synergy
HT microplate reader; BioTek Instruments Inc., Winooski, VT, USA)
with a malachite green assay (D’Angelo et al. 2001). Nitrate
(NO3-) in extracts was analyzed using colorimetric microplate
assays with a vanadium (III) chloride assay (Doane & Horwáth
2003). Ammonium (NH4+) in extracts was analyzed using colorimetric
assays with a Berthelot Reaction assay (Rhine et al. 1998).
We used the biologically based phosphorus (BBP) extraction
protocol (DeLuca et al. 2015) which uses four extractants to mimic
strategies used by plants or microbes to access P: calcium
chloride (CaCl2; dilute salt to simulate P available in soil pore
water), citric acid (C6H8O7; P sorbed to clay or weakly bound to
the soil matrix made accessible through organic acids released by
plant roots and microbes), hydrochloric acid (HCl; P strongly
bound to mineral surfaces which may be less accessible to plants
and microbes), and phosphatase (labile organic P available through
enzyme hydrolysis). Extractions were conducted in parallel by
shaking 0.5 g of each sample with 10 mL of each of the four
extractants - 0.01 M CaCl2, 0.01 M C6H8O7, 0.02 EU ml-1
phosphatase solution in a 50 mM sodium acetate buffer and 1 M HCl,
followed by centrifugation for 30 min at 3020 rev min-1. PO4 was
then determined on the supernatant from each extraction
colorimetrically, as above.
Extracellular enzyme methodology was modified from Saiya-Cork et
al. (2002) and McLaren et al. (2017). One gram of soil was blended
with buffer (modified universal buffer, pH = 7.75). Aliquots of
slurries were pipetted onto 96 well plates. Fluorescing,
4‐methylumbelliferone (MUB) tagged substrate was added to each
assay. Assays were incubated at 20 °C for 3.5 h with half‐hourly
measurements ensuring activity was measured in the linear range of
the reaction. Sample fluorescence (i.e., cleaved substrate) was
read at 360 nm excitation, 460 nm emission. For each substrate, we
measured the background fluorescence of soils and substrate and
the quenching of MUB by soils, and used standard curves of MUB to
calculate the rate of substrate hydrolysed.
Lastly, total reducing sugars methodology was modified from
Fursova et al. (2012) and Lever (1972).
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