biovolume

All perennial plants in select study plots were inventoried and measured in spring 2012. Biovolume is estimated from the maximum canopy extent along N-S and E-W axes, and plant height. For those plants (some trees, some Saguaros) that were too tall to measure with tape, height details were collected with a clinometer from level ground. The height of specimens measured by clinometer should be calculated conditionally depending on whether the value Base (deg) is positive or negative:

Base < 0, height (m) = (TAN(RADIANS(Top))+TAN(RADIANS(ABS(Base))))*distance

Base > 0, height (m) = (TAN(RADIANS(Top))-TAN(RADIANS(ABS(Base))))*distance

fertilizer_application

Fertilization with N, P, and N+P of 20 m x 20 m plots at 4 sites each in upwind, core, and downwind positions will be done twice per year to test the hypothesis that urban N deposition pushes the desert system toward P limitation. The level of N addition (as NH4NO3) is chosen to be ~ twice the current maximum deposition rate, i.e., 60 kg kg ha-1 y-1; P addition level* is 1/5 (by mass; 1/11 by atoms) that of N at 12 kg ha-1 y-1. Fertilizer is applied to treatment plots each winter and summer to coincide with the bimodal precipitation patterns characteristic of the region. The prescribed weight of dry, granular fertilizer (ammonium-nitrate for nitrogen treatments (N, N+P), triple-super phosphate for phosphorus treatments (P, N+P)) is applied using a hand spreader. As of August 2015, the rate of N fertilizer application was reduced to 30 kg ha-1 y-1, with a resulting N:P ratio of 2.5:1.

soil_ph

soil for pH analyses was collected from plant (beneath the canopy of Larrea tridentata) and inter-plant (between plant canopies) spaces within study plots. All soil within a copper ring (X diater) to a depth of 2 cm was collected with a metal spoon. The material was placed in individual plastic bags, transported to the laboratory at Arizona State University, and stored in a cool, dry location until the time of analysis. Lab procedure: (1) Mix a 2:1 ratio (by weight) of DI water and pre-sieved soil into a slurry (usually 30 mL water to 15 g soil). If the soil is highly organic, you may need to increase this ratio. (2) Wait 30 min before taking pH reading on samples. Individual samples should be started ~2 min apart to assure pH measurements are done for each sample at 30 min. During this time, calibrate the pH meter. Consult pH meter manual for calibration methods (also, see below). (3) Measure the pH of the soil slurry using a calibrated portable pH meter (be extremely careful with the tip of the probe!! Do not break it!). Press measure and wait until the pH meter has stabilized (stopped blinking) before you record the value. (4) Rinse electrode (including the bottom!) with DI water into a waste beaker between samples and dry with kim wipe but do not touch the bulb. (5) Every 10 samples check calibration in pH 7.00 buffer. If reading is off +/-0.05 or more, recalibrate the meter. (6) When finished, clean probe with DI water and place in probe storage solution (salt solution). (7) Again, if you are doing lots of soil samples, make sure that you are waiting only 30 minutes before each reading (plan so that your timing is correct by staggering addition of water by ~2 minutes).

plant_root_simulator

Soil anion and cation availability are quantified twice annually using ion-exchange membranes. Ion-exchange membranes (Plant Root Simulator (PRS) probes) are provided and analyzed by Western Ag Innovations, Saskatchewan, Canada. Probes for the analysis of soil anions and cations are deployed in pairs at locations under the canopy of Larrea tridentata (n=4) and in inter-plant spaces (n=4) within selected study plots. The probes are inserted vertically into the top 15 cm of soil, and left undisturbed until extraction after approximately 6 weeks in the ground. Upon extraction, the 8 probes of each type (anion/cation) from each habitat (plant/inter-plant) are pooled, washed with distilled water in accordance with protocols outlined by Western Ag Innovations Inc., and shipped overnight to Western Ag Innovations Inc. for analysis. PRS probe analyses typically focus on nitrogen species (i.e., NO3-N and NH4-N). However, a braoder suite of analytes is assessed approximately every 3-5 years.

stem_growth

Stem length on select Larrea shrubs are measured twice per year—once in the fall and again in the spring. Five Larrea shrubs (designated L1 thru L5) have been selected as study plants in each plot at all CN Dep sites. Measurements are taken at 4 points on each study plant—one on a stem facing each of the cardinal directions. Each time the shrubs are measured, a new piece of colored lab tape is placed around the stem, which will be used as the point from which to measure stem elongation during the following period. Field Protocol: Before arriving at the study site, cut ca. 1-inch pieces of lab tape in half length-wise and loosely stick to back of clipboard. Make enough pieces for each direction on each plant (20 per plot). Label each piece with a plant number (L1-L5) on the left side of the tape and a cardinal direction (N, S, E, W) on the right side. Study plants are marked L1 thru L5 with a silver tree tag. For each of the 5 study plants per plot: 1. Measure stem elongation: a. Look for a piece of colored tape in each cardinal direction of the shrub. Make sure the calipers are calibrated to zero and you are measuring in mm. Measure the stem (in mm) from the node directly above the piece of tape to the last node on the stem. b. If the stem branches at any point above the tape, measure each branch separately and record all the branch lengths separated by commas on the datasheet. c. Do not measure the stalks of flowers or fruits. d. If the leaves directly above or below a node have fallen off and the stem is not measurable, make a note on the datasheet. e. If the tape cannot be found, check on the ground around the shrub to see if the tape has fallen off or the branch has died. Note on the datasheet if the tape is missing or the branch is dead. Do not leave any blank spaces on the datasheet! f. Remove the old tape from the stem. 2.Measure re-taped stem lengths: a. Re-tape the stem with a new piece of tape. Place the tape close to the end of the stem, but with a measurable internode between the tape and the end of the stem. b. Measure the re-taped stem lengths as above and record on the datasheet.

leaf_tissue_icpms

Field collection: (1) Randomly choose 5 creosote plants in each of the 5 study plots for every site (15 sites total). Do not choose shrubs chosen for stem-elongation (growth) measurements. (2) From each plant, randomly select one stem in each of 4 cardinal directions. (3) From each stem, pick 4, fully developed leaf pairs from the green section of the primary stem. We are defining fully developed leaves as those that are just behind the newest leaves (the newest leaves are the 2 undeveloped green ones plus the ones just attached to them). Do not choose leaves that are farther back down the stem or leaves that are brown and/or damaged. (4) Place all leaves from each plot in a coin envelope. (5) Place coin envelopes into a dryer (60°C) upon returning to the laboratory or within 48 h. Lab processing: (1) Rinse all picked leaves with DI water in a clean fine meshed sieve and place in a coin envelope with the site name, plot name, and date processed. (2) Dry all leaves in coin envelopes at 70°C for at least 48 h before grinding. (3) Fill polycarb vial approximately 1/2 full with leaf litter, add one of the polycarb balls and cap vial, place vials in ball mill chamber and mill for 10 minutes. ICPMS: Dried and milled Larrea leaf tissue is microwave digested using a CEM MARSXpress. Approximately 0.2500g of sample was allowed to react with 10mL of Trace Metal Grade nitric acid (Fisher #A509-P212) for 15 minutes, uncapped, then ran on the CEM using the following method: 1600W at 100% power, ramped up to 180°C over the course of 15 minutes, and held at 180°C for 15 minutes. Samples were diluted at 100X prior to being run by ICP-MS. Samples were analyzed using a Thermo Fisher Scientific X Series 2 quadrapole ICP-MS and Cetac ASX- 520 autosampler. Sample introduction consisted of a conical spray chamber with impact bead and concentric nebulizer with a flow of ~1ml/min. The spray chamber was cooled to 3°C by a Peltier cooling system. Collision Cell Technology (CCT) mode was utilized to reduce interferences by the argon gas used to generate the plasma by using a mixture of 7% hydrogen and 93% helium for the following elements: Fe, Se, As, and Ca. All other elements were run in normal mode. Internal standards (Sc, Ge, Y, In, and Bi) were used to compensate for sample matrix effects and instrument variability.

annuals_biomass

Each spring when there is sufficient rainfall, we estimate aboveground net primary productivity (ANPP) of herbaceous annual plants in each of the Desert Fertilization Experiment study plots by harvesting the aboveground portion of annual herbs from a 1/2 m x 1/2 m section of established 1 m x 1 m sub-plots. These plots are distinguished from similar 1 m x 1 m sub-plots that are used for community composition. Take care to avoid the community composition sub-plots! Only harvest the sub-plots that are designated for biomass. Protocol overview: In each Desert Fertilization experiment plot, we established 4 biomass sub-plots per plot, two in each of two patch types, between shrubs (IP for inter-plant space) and under the canopies of Larrea tridentata (P for under plant). Each sub-plot is coded with the patch type (IP or P); the letter B for biomass (to distinguish it from the community composition sub-plots, which are not harvested and do not have a letter B after the patch type code); and the sub-plot number (1 or 2). For example, IPB1 & IPB2 are interplant space sub=plots one and two, and PB1 & PB2 are sub-plots 1 and 2 under Larrea tridentata shrubs. Remember that the biomass sub-plots are distinguished from the community composition sub-plots with the letter B after the patch type code (example, IPB1 = Biomass sub-plot 1, inter-plant space). Within each 1 m x 1 m biomass sub-plot, teams will clip aboveground annual biomass from one 1/2 m x 1/2 m quadrat in each 1-meter biomass sub-plot. This sub-section is rotated each year so that any given patch of ground is harvested at the most once every four years. Teams will carefully write the plot identifier on paper bags. (Double-check the labels! Labeling mistakes are the most common error in this procedure). Then teams will cut all green and brown herbaceous plant biomass that occurs within the boundaries of the 1/2 m x 1/2 m sub section and place it in the pre-labeled bag. After the harvest is complete, teams will dry the bags and biomass at 60°C for 48 hr then weigh the biomass (subtracting the weight of the bag) on a scale and record.

annuals_composition

Overview: In Spring 2008 within each site, the CAP LTER randomly chose and permanently marked four 1 m x 1 m sub-plots within each 20 m x 20 m main plot, split between two patch types, under L. tridentata shrubs and inter-plant spaces (hereafter, under L. tridentata, P and Inter-plant space; IP). At peak herbaceous biomass (in approximately March of each year), percent cover of all annual herbaceous species rooted within these four plots (two each under L. tridentata and in interplant spaces i.e. between shrubs) will be estimated. Note that there are also four 1 m x 1 m Biomass sub-plots in each 20 m x 20 m plot – these are different than the community composition plots described here because they are clipped each year while the community composition plots are not. Care should be taken in the field to identify the correct four plots for community composition and avoid the four plots to be harvested for biomass. Steps: 1. On the top of the data sheet (See Table 3 for sample), record the date; names of the sampling crew (i.e. plant identifier, recorder); site (three letter code); plot number (1 to 75); treatment (C1 – no sampling from C2 plots, Nitrogen [N], Phosphorus [P], and Nitrogen+Phosphorus [NP]). Double check the plot numbers and sub-plot numbers! 2. All observers should first calibrate their percent cover estimates on several test plots in the field prior to percent cover measurements. You want to make sure all botanists are seeing the plots similarly. 3. Place the 1 m x 1 m quadrat over the corner rebar at each 1-m2 plot. Working in teams of two (one recorder, one plant identifier), proceed through the steps below. 4. Estimate and record (at the bottom of the data sheet) the relative areal cover of plant matter (e.g. green material) versus bare ground (e.g. brown material) at the soil surface in the plot (e.g. 70% green, 30% brown). If there are soil crusts, measure those as well in a separate category. 5. Identify which annual species are present in the plot and mark a small dot in the corner of the datasheet to easily find later. Use roughly the same amount of effort in each plot to find plant species. As you get better, it will take less time because you are getting more skilled; as you get tired, it will take more time; in simpler systems, it will take less time. There is no a priori way to know when you are using the same amount of effort, but this is usually approximated by time (in minutes: typically 1-3 minutes per species). Each botanist should do some test plots to see how long it takes him/her to find all the species in the plot. Different botanists will take different amounts of time to apply the same effort– this is ok – but the same botanist should use approximately the same amount of time per plot. 6. Next, estimate and record the cover of each species individually to the nearest whole number (e.g., 1%, 2%, 3%...97%, 98%, 99%, 100%). If a species is present in a plot, but does not cover a full 1%, record it as <1%. The sum of all individual species cover can be greater than 100% since some species overlap. 7. Finally, estimate and record the cover of the overstory shrubs within the plot or overhanging the plot. We estimate cover for all annuals rooted in the quadrat, but areal cover of any shrubs rooted in or hanging over the plot. For IP plots, this will be zero most of the time (i.e. unless there is a large shrub nearby that has grown over the plot). For P plots, this will be have two components: (1) the areal projection of the shrub onto the 2-D grid, and (2) the area of the stem base or any other shrub stems. It is expected that the cover of the shrubs overlap the herbaceous species. However if two shrub species overlap in their cover aggregate these into a single larger individual (rare case). 8. Unknown species are recorded with unique identifiers (e.g., Unknown 1, Unknown 2, etc) along with a visual description of the plant and a picture to be used to help identify the unknown in the lab. When possible, a sample of the unknown species should be collected from outside of the plot in a labeled coin envelope and brought to the herbarium to be identified. Unknown species are usually very small and_or undeveloped, and may not be able to be identified to species – this is ok. Err on the side of lumping unknown species into other known species in the same plot. 9. Use caution not to include the percent cover of desiccated/dead plants within the plots from the previous season (these can usually be identified easily as white, old plant material). However, depending on the timing of data collection, some herbaceous plants from the current growing season may have passed peak biomass and already begun to desiccate (dry out or senesce). In this case, estimate the percent cover of these plants as if they were at peak biomass. 10. Before you leave a site, confirm that you have datasheets for all plots and sub-plots, and that there are no duplicates or ambiguous plot labels on your sheet.

annuals_composition_2008

In the spring of 2008, the estimation of the cover of annuals in the study plots was conducted uniquely relative to subsequent years: collector SLC allowed for layering within the herbaceous layer so that total cover when you summed across species could be > 100% (potentially limitless), whereas collector CMC did not allow for layering within the herbaceous layer such that the cumulative sum of cover of annuals cannot be more than 100%. To facilitate comparison, data collected by SLC were relatavized by the total annual cover in the study plot (i.e., the percent of the plot covered by annuals) and the cumulative sum of the cover of annual species in the study plot (i.e., % cover of plant * (Total Plant Cover in Plot / sum of all species (sumSpp))) such that the estimates by both collectors reflect the relative cover of each plant taxon. In subsequent years, values reflect a collection method that allows for layering. Owing to the requisite relativization, data collected by SLC in 2008 were reconstructed from original field data sheets. As such, information gained from post-processing that resolved the identity of species that were not identified in the field was not available resulting in a greater number of unidentified species (unknowns and variants) for data collected by SLC relative to CMC.