In 2005, annually harvested root ingrowth donut structures were co-located with previously established mini-rhizotron tubes established at four sites on McKenzie Flats located on the east side of Sevilleta NWR: 10 replicate structures in both burned and unburned blue and black grama dominated grassland plots at Deep Well, 10 replicates each on nitrogen fertilization plots and respective control plots on McKenzie Flats(20 total), 10 replicates in creosote dominated shrubland at the Five Points Creosote Core site and in 2011, 13 structures were put in the Monsoon site. Roots and soil are harvested annually in late fall after the growing season, and structures are reestablished in situ for consecutive harvests each year. Each structure allows roots to be harvested at two depths (0-15 and 15-30 cm) to estimate root production, or below ground net primary productivity. In order to compare estimates of root production from two methods, root ingrowth donuts were collocated with mini-rhizotron tubes at all localities except for the burned grassland plot at Deep Well.
Experimental Design Annually harvested root ingrowth donut structures are co-located with previously established mini-rhizotron tubes established at four sites on the McKenzie Flats on the east side of Sevilleta NWR: 10 replicate structures in both burned and unburned blue and black grama dominated grassland plots at Deep Well, 10 replicates each on nitrogen fertilization plots and respective control plots on McKenzie Flats (20 total), and 10 replicates in creosote dominated shrubland at the Five Points Creosote Core site. Methods were adapted from Milchunas et al (2005). We use a formidable 10 diameter soil core to create cylindrical holes in the ground to a depth of 30cm without disturbing soil profile at the cylinder walls. The soil core was inserted with a slide hammer and had to be removed each time with a come-along mounted on a steel-pipe tripod. Walls were subsequently lined with plastic cross-stitch craft work canvas (macram mesh) which supports cylinder walls through time but allows roots to pass through. Two pieces of 6 diameter PVC were placed in the center of the larger cylindrical hole, set in place with bags filled with sand that act as ballasts. The two pieces of PVC were beveled on opposite ends to fit together and prevent movement of the donut center. The top cylinder went to a depth of 15 cm and the bottom piece went to a depth of 30 cm, representing 0-15 and 15-30 cm in the soil profile when stacked upon one another. Finally, sifted soil from the location was used to fill the space between the plastic canvas lining the hole wall and the PVC pipe placed in the center. It is this soil which is harvested annually at two depths. A PVC cap was placed on top of the PVC to eliminate water infiltration from rain through the donut center and to keep sunlight from disintegrating the sand bag ballast. All root ingrowth donuts were GPSed.
Sample Harvest Root donuts are harvested annually in November after the growing season. Roots are harvested by first removing the sand bags from the top cylinder and placing a bowl into the center of the cylinder. The top cylinder is then removed. Soil and roots are cut away from the cylinder wall and collected. This harvest procedure is then repeated for the lower half of the donut structure. Soil and roots collected are placed in a separate plastic bag for each depth. Once the soil and roots are harvested, the root ingrowth structure is rebuilt. After harvest, soil and root samples are stored in a chest freezer until they can be processed.
Sample Processing The total volume of soil from each sample is measured and recorded. Soils are filtered through a series of sieves in which to harvest the roots present in the sample. The roots are then repeatedly rinsed to remove all the soil from the sample, dried at 60 degrees C, and then weighed.
23 Jan 2009All data sets (2005-2008) were combined and checked for errors in excel and exported into Navicat. From the 2007 data, I converted the dry root mass from grams to milligrams and changed depth data to be 0-15 and 15-30 cm. QA/QC'd data. I deleted data line from DWB sample 7, depth 15-30 cm with volume 2600 ml because it was a duplicate. I also changed the depth of DWB sample 12, depth 15-30 cm with volume 2000 to the depth 0-15 cm because the depth 15-30 cm was duplicated. -Changed missing data on volume and weight due to plant being dead to -888. -Changed missing data on volume and weight due to human error to -999. --A. Swann
Filtered data in Excel then exported it into Navicat using the import wizard.
Additional Information on the personnel associated with the Data Collection / Data Processing
Sevilleta Field Crew Employee History
Chandra Tucker April 2014-present, Megan McClung, April 2013-present, Stephanie Baker, October 2010-Present, John Mulhouse, August 2009-June 2013, Amaris Swann, August 25, 2008-January 2013, Maya Kapoor, August 9, 2003-January 21, 2005 and April 2010-March 2011, Terri Koontz, February 2000-August 2003 and August 2006-August 2010, Yang Xia, January 31, 2005-April 2009, Karen Wetherill, February 7, 2000-August 2009, Michell Thomey, September 3, 2005-August 2008, Jay McLeod, January 2006-August 2006, Charity Hall, January 31, 2005-January 3, 2006, Tessa Edelen, August 15, 2004-August 15, 2005, Seth Munson, September 9, 2002-June 2004, Caleb Hickman, September 9, 2002-November 15, 2004, Heather Simpson, August 2000-August 2002, Chris Roberts, September 2001-August 2002, Mike Friggens, 1999-September 2001, Shana Penington, February 2000-August 2000.
We studied the diversity of arbuscular mycorrhizal fungi (AMF) in a semiarid grassland and the effect of long-term nitrogen (N) fertilization on this fungal community. Root samples of Bouteloua gracilis were collected at the Sevilleta National Wildlife Refuge (New Mexico, USA) from control and N-amended plots that have been fertilized since 1995. Small subunit rDNA was amplified using AMF specific primers NS31 and AM1. The diversity of AMF was low in comparison with other ecosystems, only seven operational taxonomic units (OTU) were found in B. gracilis and all belong to the genus Glomus. The dominant OTU was closely related to the ubiquitous G. intraradices/G. fasciculatum group. N-amended plots showed a reduction in the abundance of the dominant OTU and an increase in AMF diversity. The greater AMF diversity in roots from N-amended plots may have been the result of displacement of the dominant OTU, which facilitated detection of uncommon AMF. The long-term implications of AMF responses to N enrichment for plant carbon allocation and plant community structure remain unclear.
Sampling Design Roots from three Bouteloua gracilis plants were collected from three control and three N-amended plots (a total of 72 plants).
Field Methods Plant samples were collected in plastic bags, kept at 4 C and processed the same or next day after collection.
Lab Procedures Whole roots from one plant from each plot (6 plants/date) were cleaned under running tap water, rinsed twice with sterile water, and dried on paper towels. Roots were determined to be alive if they did not exhibit lesions, were not obviously damaged, possessed a prominent number of root hairs and were connected to green leaves. A subset of these roots were microscopically analyzed to confirm AMF colonization, and the others were stored at -20 C until their DNA was extracted.
Microscopy The microscopy results showed very low AMF colonization, as a compromise between the need for a large sample of clones (to reliably characterize the AMF community within each plant) and a significant sample of plants from control and N-amended plots (to study the effect of nitrogen on colonization), we decided to select 3 plants from control and 3 plants from N-amended plots and sequence approximately 100 clones per plant using fungal and AMF specific primers.
DNA Extraction Three to five roots segments (approximately 3 cm long) from each plant were used for DNA extraction. DNA was extracted using a DNeasy plant Mini Kit (Qiagen, Chatsworth, CA). Primers NS31 and AM1 were used to specifically amplify AMF (Helgalson et al., 1998; Simon et al., 1992). For each of the 6 root samples, a total of 25 to 47 random clones were sequenced, 260 sequences in all. PCR was performed using the following protocol: initial denaturation at 95 C for 5 min, followed by 30 cycles of 95 C for 30 s, 53 C for 30 s, and 72 C for 45 s, with a final extension of 72 C for 7 min. DNA was amplified in 25 uL reactions with 12.5 uL Premix Taq (Takara Bio), 1.0 uL of each primer (5 uM), 3 uL of BSA 1%, 6.5 uL of milliQ water, and 1 uL of template DNA. The first PCR products were cleaned with ExoSAP-IT (USB, Cleveland, Ohio) and 1 uL of the cleaned PCR product was used as template for the second PCR. Products were cloned with TOPO-TA cloning kit (Invitrogen, Carlsbad, CA) following the manufacturer's instructions. Clones were amplified and sequenced using rolling circle amplification (TempliPhi, Amersham, Buckinghamshire, England) and BigDye Terminator v1.1 Sequencing Kit (Applied Biosystems, Foster City, CA), respectively. Sequencing was conducted at the Molecular Biology Facility of The University of New Mexico. Forward and reverse sequences were assembled and edited with Sequencher 4.0 (Gene Codes, Ann Arbor, MI).
Sequence Analysis The program CHIMERA CHECK 2.7 of the Ribosomal Database Project (http:// rdp.cme.msu.edu/html/analyses.html) was used to check for chimeric 18S nrDNA sequences. Sequences were BLASTed against GenBank and information from GenBank obtained using phd, bioperl scripts and a mysql database written by George Rosenberg, Molecular Biology Facility of the University of New Mexico. Glomeromycota sequences were submitted in GenBank under accession numbers EF154520 and EF154698. OTUs were determined using the DOTUR program (Schloss and Handelsman, 2005). Distance matrices generated with the F84 evolutionary model using the DNADIST program from PHYLIP (Felsenstein, 2005) were used as input files to DOTUR. A similarity level of 97% has been used as the lower boundary to define OTUs in several studies of AMF (e.g. Helgason et al., 1999). We performed our analysis using both 97 and 99% of similarity to evaluate how mycorrhizal fungi at different taxonomic levels respond to N deposition. Rarefaction curves and diversity estimators (Chao, ACE) were calculated for the pooled data (N and control plots) and for N and control treatments with DOTUR. The effect of N enrichment on the AMF community also was evaluated in a phylogenetic context using UniFrac (Lozupone and Knight, 2005). The UniFrac metric estimates differences between microbial communities inhabiting different environments based on phylogenetic distances. In our study, we used this metric to evaluate the percentage of branch length in a phylogenetic tree that leads to descendants from N- amended and control plots. A phylogenetic tree generated with PAUP 4.0b10 (Swofford, 2002) that include all the Glomeromycota sequences from this study was used as input file to calculate UniFrac significance. Trees were constructed using the neighbor-joining (NJ) algorithm and maximum parsimony (MP) in PAUP 4.0b10 (Swofford, 2002). Bootstrap values were estimated from 1000 replicates for the MP and NJ analysis. A NJ phylogenetic tree that includes only representative OTUs (defined at 99% similarity with DOTUR) was used for normalized weighted principal coordinate analysis in UniFrac (Lozupone and Knight, 2005). The weighted UniFrac accounts for the relative sequence abundance in each sample. The NJ tree and a text file that includes OTU abundances for each sample were used as input files.
PTC 200, Pertier Thermal Cycler PCR machine DNA Engine
Changes to the data: File created on 6/13/2008 by Andrea Porras-Alfaro.
Data are available from Genbank: http://www.ncbi.nlm.nih.gov/Entrez/index.html Accessions EF154520-EF154698
Maintaining high rates of water loss during times of high resource availability could allow establishing woody desert perennials to grow quickly by allowing them to take advantage of the fleeting but abundant monsoonal moisture typical of warm deserts like the Chihuahuan. However, a plant cannot endlessly increase water loss in order to grow faster --there are hydraulic constraints on rates of water loss. The hydraulic properties of each particular plant xylem and soil microsite, as well as the AR:AL absorbing root area to transpiring leaf area ratio) interact to set limits on rates of water loss. If transpiration rates become too high, cavitation may limit the ability of the xylem to supply water to the leaves. The main objective of this study was to test two hypotheses on a population of Larrea tridentata at the Sevilleta LTER in central New Mexico (1) do small plants grow faster and use water less conservatively than large, and (2) are there differences in the hydraulic constraints on small and large plants. Measurements were made every six weeks in the spring, summer and fall from April 2002 - August 2003. Field measurements of shoot growth, gas exchange and plant and soil water potentials were made to determine growth rates and water use. Measurements of leaf specific conductance determined the ability of the xylem to supply water to the leaves. Excavation findings were used to estimate (AR:AL). Xylem vulnerability curves and soil texture analysis were used to determine the hydraulic properties of the plant xylem and soil. A model determined where the limiting conductance occurred in the plant-soil continuum.
For gas exchange measurements a LiCor 6400 portable gas exchange system was used. Measurements took place in June, August and September 2002 and May, June, and August 2003. Two measurements were made on each plant at approximately 7-9:30AM and 10-12:30PM. One branch tip was chosen and marked on each 10 small and 10 large plants. The same branch tip was used for measurement throughout the day unless it broke, at which time another branch tip was chosen and marked. Stomatal Ratio was set to one because stomates are present on both sides of the leaf in this species. Because of the small size of the leaves, an energy balance approach was used to calculate the leaf temperature in the chamber.
Chamber temperature and humidity were controlled at ambient and reference CO2 was set to 400ppm. Using natural light plants were clamped into the chamber, oriented in their original direction, chamber conditions were allowed to stabilize. Leaf area was set to one during measurement. Because of the small leaves of the species, a branch tip had to be measured. Measured branch tips were cut and returned to the lab where their leaf area was measured.
A Vista Scan Scanner was used to create an image of the leaves. The bitmap image was then analyzed for number of pixels using Scion Image. A regression equation was developed which converted pixel number into leaf area in cm2.The gas exchange data was then recalculated to adjust for leaf area.
For plant water potential a Scholander Pressure bomb was used to measure branched stem tips consisting of 15-20 leaves and a woody base. Predawn water potential samples were collected between 4AM and 5AM. Midday water potentials were collected between 11AM and 1PM. Samples were placed into a plastic baggie with a moist paper towel during transport. Samples were collected in May, June, August, September and December 2002 as well as January, March, May, June and August 2003.
For whole plant hydraulic conductance measurements large sections of the xylem were measured using a vacuum canister to generate known vacuum pressures. The plant was attached to a water filled container on a balance via tiagon tubing. Changes in weight on the balance, and thus flow rate (mg/s) through the plant were measured by a computer using a program written in Turbo Pascal. Each sample was measured at four or five pressures and the change in flow rate with pressure was calculated as the total hydraulic conductance of all tissues contained in the sample. All samples were immediately placed into a plastic bag with a wet paper towel and transported to the lab where they were measured within 48 hours of collection.
For large plants an entire stem of the plant was cut at the base. The stems were cut under water at the lab, to about 30cm. For small plants the entire plant was excavated, and any roots larger than about 2mm in diameter were kept intact. Back at the lab, most of the root system was cut off under water, leaving the root collar and the initial un-branched portion of the main root which obviously supplied the entire plant. For both sizes, all green material was removed from the tips of the branches, leaving only woody stems.
The green portion of the plant included all leaves and sometimes large (up to 150mm) branched sections. For root hydraulic conductance measurements root segments +AD4-50cm were cut in the field and transported to the lab wrapped tightly in 3 plastic bags containing wet paper towel. Segments were re-cut underwater and the ends shaved off with a razor blade. They were then placed on a manifold and flow through the segment was measured. Segments were then flushed for 15 minutes with distilled water at 100kPa. Flow was then re-measured. Percent loss of conductivity is calculated as the difference between pre and post flush flow divided by post flush flow and multiplied by 100. Due to time constraints only one flush was performed on each sample. For soil water potential monitoring soil thermocouple psychrometers were placed under 4 large and 4 small plants at 30cm and 45cm below the soil surface. Measurements were made at or around 2AM when temperature gradients were at a minimum. A Campbell datalogger reported millivolt output which was converted to MPa using calibrations determined in the lab. Calibration involved regression of millivolt output against solutions of known salt concentration for each psychrometer.
Additional Information on the personnel associated with the Data Collection / Data Processing Joy Francis, a post-doc with Jim Gosz, was instrumental in setting up this study.