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).
References cited
D’Angelo E, Crutchfield J, Vandiviere M., (2001). Rapid,
sensitive, microscale determination of phosphate in water and
soil. Journal of Environmental Quality 30:2206-2209.
DeLuca, T. H., H. C. Glanville, M. Harris, B. A. Emmett, M. R. A.
Pingree, L. L. de Sosa, C. Morenà, and D. L. Jones. (2015) A novel
biologically-based approach to evaluating soil phosphorus
availability across complex landscapes. Soil Biology and
Biochemistry 88:110-119.
Findlay, R.H., (2004). Determination of microbial community
structure using phospholipid fatty acid profiles, in: Molecular
Microbial Ecology Manual, 2nd Edition. Kluwer Academic Publishers,
Netherlands, pp. 983-1004.
Fursova, O., Pogorelko, P., Zabotina, O.A., (2012). An efficient
method for transient gene expression in monocots applied to modify
the Brachypodium distachyon cell wall, Annals of Botany, Volume
110, Issue 1, pp. 47-56, https://doi.org/10.1093/aob/mcs103.
Doane, T.M., Horwáth, W.R., (2003). Spectrophotometric
Determination of Nitrate with a Single Reagent. Analytical Letters
36(12), pp. 2713-2722.
Lever, M., (1972). A new reaction for colorimetric determination
of carbohydrates. Analytical biochemistry, 47(1), pp.273-279.
McLaren JR, Buckeridge KM, Van de Weg MJ, Shaver GR, Schimel JP,
Gough L (2017) Shrub encroachment in Arctic tundra: Betula nana
effects on above- and belowground litter decomposition. Ecology
98:1361-1376.
Rhine, E.D., Mulvaney, R.L., Pratt, E.J., Sims, G.K. (1998).
Improving the Berthelot Reaction for Determining Ammonium in Soil
Extracts and Water. Soil Science Society of America Journal,
62(2), 473-480.
Saiya-Cork KR, Sinsabaugh RL, Zak DR (2002) The effects of long
term nitrogen deposition on extracellular enzyme activity in an
Acer saccharum forest soil. Soil Biology and Biochemistry
34:1309-1315
U.S. Salinity Laboratory 1954. Diagnosis and improvement of saline
and alkali soils. U.S. Dept. of Agric. Handb. 60. U.S. Gov. Print.
Office, Washington, DC.
