METHODS
This work builds on previous research at Idaho State University’s Barton Ecological Research Area in Pocatello, Idaho. For detailed descriptions of the original experiment, see:
Inouye, R. S. (2006). Effects of shrub removal and nitrogen addition on soil moisture in sagebrush steppe. Journal of Arid Environments 65:604-618. doi: 10.1016/j.jaridenv.2005.10.005
Bechtold, H. A., & Inouye, R. S. (2007). Distribution of carbon and nitrogen in sagebrush steppe after six years of nitrogen addition and shrub removal. Journal of Arid Environments 71:122-132. doi: 10.1016/j.jaridenv.2007.02.004
In brief, the original experiment was initiated in 1997 to evaluate the ecological effects of shrub removal and nitrogen (N) addition in a sagebrush steppe ecosystem (42.853 N, 112.402 W, 1450 m elevation). Thirteen plots (20 x 20 m) were established in a 1.4 ha area and randomly assigned to treatments. Three plots were assigned to shrub removal treatment, in which shrubs were cut at ground level and removed. Three plots were assigned to low N addition (6 kg N/ha/yr), and another three plots were high N addition (12 kg N/ha/yr), using hand broadcast ammonium nitrate. The remaining four plots were unmanipulated controls. These experimental conditions were maintained annually until 2010, after which they were no longer maintained.
On August 15, 2020, a wildfire burned the entire area encompassing the original experiment. Shortly after, we began a study to evaluate how legacies of prior shrub removal and N additions in the original experiment affect post-fire plant recovery. First, we added two new 20 x 20 m plots as controls to enhance statistical power to the original experiment. Then we modified the experiment to evaluate the effects of native seed additions intended to enhance biotic resistance against invasive plants. To study seed additions, we split each of the 20 x 20 m plots into four new plots (10 x 10 m) and randomly assigned each set of new plots to four seed treatments. On April 5, 2021, three seed mixes were hand sown at a rate of 0.2 g/m² of pure viable seed (purchased from Western Native Seed, Coaldale, Colorado). The grass-only mix consisted of two species: 50% Elymus elymoides and 50% Koeleria macrantha. The forb-only mix consisted of four species: 25% Helianthus petiolaris, 25% Oxytropis sericea, 25% Rudbeckia hirta, and 25% Sphaeralcea coccinea. The third mix included the same six species of grasses and forbs, but had a composition of 25% per grass species and 12.5% per forb species (by pure viable seed). The fourth seed treatment was a control without experimental seed additions. These seed treatments were in addition to the application of a base seed mix of 16 grass, forb, and shrub species that was hand sown over the entire burned area at a rate of 1.4 g/m² (purchased from Idaho Grimm Growers Warehouse Corp., Blackfoot, Idaho). Species used in the experimental treatments were not included in the base mix.
Within each new plot (10 x 10 m), we established four 1 m² permanent quadrats for data collection activities. At a microhabitat scale, shrubs may alter the environment in ways that have lasting effects after they have died, such as legacy effects on soils. To understand how these legacy effects interact with those of shrub removal and N additions, we contrasted areas with and without former shrubs within the experimental plots. A flag was haphazardly tossed within the plot, then a quadrat was established adjacent to the nearest shrub stump, and another quadrat was established at least 1 m away in an area between shrub stumps. This process was repeated for the next two quadrats. In total, there were 240 quadrats distributed across 60 plots.
In addition, we established auxiliary experiments to better understand conditions in the primary experiment. Part of the field site did not burn in the wildfire, which allowed us to contrast burned vs. unburned conditions. We established five new plots (10 x 10 m) in the unburned area, with four 1 m² permanent quadrats for data collection activities in each plot. The unburned plots did not receive base or experimental seed additions, and to evaluate the effect of fire independent of seed additions, we established another five new unseeded plots (10 x 10 m) and associated sampling quadrats in the burned area to compare with the unburned area. Furthermore, we conducted an auxiliary study to evaluate the effects of seed density for the seed addition treatments. In each of the control plots from the original experiment (without shrub removal or N additions), we established three more 1 m² permanent quadrats within the plots receiving grass, forb, or grass/forb seed treatments (quadrats placed irrespective of former shrub location). One quadrat received seed at half the rate sown into the surrounding plot, another received two times this rate, and the last received four times this rate. Lastly, within the same plots we established a fourth 1 m² quadrat to assess the effects of seed application timing. Following the collection of the present dataset, all seed additions were repeated in the experimental plots on November 22, 2021, except for the group of quadrats reserved to evaluate the effects of seeding timing. Instead, this treatment group received a second application of seeds on April 21, 2022. Across the three auxiliary studies, we established a total of 112 quadrats in addition to those created for the primary experiment (352 quadrats across all experiments).
In Summer 2021, we measured plant variables on July 7-8. Within each quadrat, we measured cover for each plant species and height of the tallest vegetation. Adjacent to each quadrat, we collected and weighed aboveground biomass. We measured soil variables on July 22-23 and 28-29, 2021, with samples collected along the edge of the quadrats. Soil data collection was focused on the 280 quadrats in the primary experiment and the burned-unburned auxiliary experiment. For each of these quadrats, we measured soil moisture, pH, electrical conductivity, ammonium-N content, nitrate-N content, and phosphate content. We also measured N mineralization using in situ incubation. Nutrient contents were measured on a Discrete Analyzer in the Lohse Lab. Remaining soil samples were archived for later analysis. See the protocols provided below for details of these procedures.
EXPERIMENTAL PROTOCOLS
List of experimental protocols used in this study, which follow in the below order:
1. Plant Surveys
2. Aboveground Biomass
3. Soil Collection
4. Fresh (Field Condition) Soil Sieving
5. Gravimetric Water Content
6. Nutrient (NH4, NO3, PO4) Extractions
7. pH & Electrical Conductivity
8. Archiving Soils for Later Analysis
9. In Situ Nitrogen Mineralization
1. PLANT SURVEYS
Materials
● PVC quadrat with markings every 0.1 m on each edge
● Long piece of PVC with markings every 0.1 m
● Flag
● Plant survey datasheets
● Clipboard
● Plant ID guide
Procedure
1. Record the USDA code of every plant species present in the quadrat
2. Identify the tallest plant rooted in the quadrat, record USDA code. Using a meter stick, measure the height to the nearest 0.1 cm (without disturbing or straightening the plant).
3. Lay the PVC quadrat with markings over the established quadrat markings. Line up the piece of PVC with the first 0.1 m mark on either edge of the PVC quadrat.
4. Starting at the first 0.1 m mark on the PVC piece, drop the flag (or pin) without looking. Record a tick mark next to the plant USDA code that is touching the pin at the highest point. The pin may also hit bare ground, plant litter (dead plants, herbaceous material or woody i.e. stumps), rock, or fungi.
5. Repeat with the next 0.1 m mark on the piece of PVC. Once this row has been completed, move the piece of PVC up to the next 0.1 m mark on the PVC quadrat and repeat steps 2-3.
6. Ensure that recorded tallies add up to 100 for each quadrat surveyed.
NOTE: There are 100 pin counts for each quadrat, totaling 100% cover. There are markings every 0.1 m, but not directly along the edges because each pin point is at the center of a 1 dm² area. To better understand the contribution of species that were present, but not recorded as a pin count, we added one to each species’ pin count in the dataset to record their presence. We repeated this method to record rock and fungi presence. We did not add one to the pin counts of bare ground and litter in the dataset.
2. ABOVEGROUND BIOMASS
Materials
● Pre-labeled brown paper bags
● Extra paper bags
● Sharpie
● Tape
● Clippers
● 1/16 m PVC quadrat
Procedure
1. Lay the 1/16 m PVC quadrat in the corner that is being sampled (NE corner in summer 2021).
2. Cut all plants present in the quadrat at ground level using clippers and place in a paper bag. Tape shut, ensuring the label is still visible. If necessary, label and use an additional paper bag for plant material.
3. Store plant samples at room temperature, ideally in a hood.
NOTE: Only plants rooted in the quadrat were considered for aboveground biomass in 2021.
3. SOIL COLLECTION
Materials
● Pre-weighed and labeled gallon Ziploc bag for corresponding quadrat
● Soil corer + hammer (AMS Hammer-Head Soil Probe Kit, Gemplers Item#212133)
● Cooler + ice packs
● Vinyl or nitrile gloves
Procedure
1. Using a ruler, mark 10 cm on the soil corer with a sharpie. This is the depth of the soil core that will be taken.
2. Before entering the plot, drive the corer into the ground outside to give the corer a “dirt bath”. to sample the quadrat, line the corer on the middle of one edge of the quadrat, just outside of the quadrat boundaries.
3. Keeping the corer vertical, drive it into the ground by hammering the mallet on top of the corer until the 10 cm sharpie mark is flush with the soil surface.
4. Carefully lift the corer out of the ground, being careful not to lose any material. Place the tip of the core into the pre-labeled Ziploc bag. Shake the core into the bag or use a gloved hand to push the core into the bag. This step is easiest if one person holds the bag open while the other person transfers the core into the bag.
5. On the same edge, take another core adjacent to the first samples using steps 3-4. Transfer the second core into the same bag with the first core.
6. Repeat steps 3-5 on the opposite side of the quadrat. All four cores should be placed into the bag, so each quadrat has one bag. Before closing the bag, push all of the air out and ensure a tight seal. Four cores are necessary to ensure enough mass for all soil processing.
7. Place the sample bag into a backpack and move onto the next quadrat. After sampling all quadrats in a plot, change gloves.
NOTE: For sampling in Summer 2021, two soil cores were taken from the W and E outer edge of each quadrat (four cores total). In teams of two, one person was the soil corer and the other would lead the team to the correct quadrat to sample next, keep track of bags, and hold the bag for the corer. Partners would switch jobs every couple of plots. After each plot was sampled, the bags of soil were transferred from the backpack to the cooler. Nutrients needed to be extracted immediately after soil collection; therefore, the team would collect soils in the morning, and return to the lab in the afternoon to process soils.
4. FRESH (FIELD CONDITION) SOIL SIEVING
Introduction
Mineral soil consists of sand, silt, and clay particles that all measure less than 2 mm in diameter. To ensure that you are analyzing only mineral soil, field collected soils are sieved. Sieving also helps homogenize the soil by breaking up aggregates that can store soil organic matter.
Materials
● 2 mm (No. 10) stainless steel sieve and bottom
● Wire brush
● Paper towels
● 70% EtOH or isopropyl alcohol
● Weighed and labeled Ziploc bags (quart-size for coarse fraction)
● Forceps
● Balance
Procedure
1. Bring fresh field soil back to the lab and lay bags on the countertop.
2. Before sieving the soil, dampen a clean paper towel with 70% EtOH. Wipe out the base and mesh parts of the sieve. Run another dry paper towel over the sieve to remove all visible signs of dirt and dust. Ensure the sieve is completely dry and that there is no trace of EtOH before pouring in the soil.
3. Place the mesh on top of the base. Carefully pour the entire bag of soil onto the mesh. Gently shake and tap the sieve to get all soil possible through the sieve.
4. Using a gloved hand, rub the soil through the mesh. If needed, use the wire brush to move the soil through the sieve. Do not push too hard on the mesh, this can damage the sieve.
5. Ensure that all of the soil has gone through the mesh by gently running your hand over the mesh surface and ensuring all aggregates are broken up. Barton road soils are dry, so there may be small aggregates of soil that look like rocks.
6. Pick any plant material out of the mesh part of the sieve and discard. Transfer rocks (coarse fraction >2 mm) into a tared weigh boat and record weight into the datasheet. Discard rocks after ensuring weight was recorded.
7. Pick any large roots (>2 mm in length) out of the sieved soil using forceps. This is a coarse root picking and should not take too much time.
8. Pour the sieved soil (< 2 mm) into the original pre-weighed Ziploc bag.
9. Weigh soil in Ziploc bag.
NOTE: This is the first thing that needs to be accomplished once soils are brought back from the field. Ideally, there would be a team of 6-8 personnel present to process the soils in a timely manner. Immediately upon return to the lab, as many people as possible should start sieving. After about half the soils are sieved, one person or a couple (depending on how many people present) can switch to weighing out soils for gross water content measurements and KCl extractions. Barton soils have few rocks, so it is unlikely that there will be much of a coarse fraction.
5. GRAVIMETRIC WATER CONTENT
Introduction
Gravimetric water content (GWC) is often determined anytime other soil properties are measured. We must measure GWC in conjunction with nutrient extractions to determine the amount of dry grams of soil.
Materials
● Soil tins with label tape (Forestry Suppliers #77043, tape: Scotch Model#2020-24AP)
● Spoon
● KimWipes and isopropyl alcohol to clean spoon between samples
● Desiccator and fresh desiccant
● Coarse balance (0.01 g)
● 105 C oven
Procedure
Once soils have been sieved, subsample soil for gravimetric water content into a tin:
1. Place a piece of tape on the lid of a soil tin. Label with the sample ID.
2. Tare coarse balance and weigh the labeled soil tin with the lid on. Record weight into spreadsheet (tin weight).
3. Place 20-30 g (three spoonfuls) of sieved (<2 mm) soil into the labeled tin. Weigh and record (wet soil weight + tin weight).
4. Place soil tins (with lid cracked, but still on top of base) in a 105 C oven for 24 to 48 hours.
5. After at least 24 hours, weigh tins. Working in batches of ten, remove the tins from the oven with a gloved hand. Weigh and record (dry soil weight + tin weight) immediately.
6. Once all tins have been weighed and recorded, dump tins into trash and remove tape.
7. Calculate the soil moisture or gravimetric water content of soils:
Soil moisture (GWC) = ((soil wet weight - tin weight) - (soil dry weight - tin weight))/(soil dry weight - tin weight)
NOTE: On processing days with 6-8 people, 2-3 people could begin subsampling already sieved soils into tins and specimen cups while the rest of the team finished sieving all soils. Ideally, two people would each be sitting in front of a scale. The third person would label soils tins with the soil ID (from sieved soil gallon bag) and place the tin and bag of soil next to the person at the scale. The labeler would continue this process with soil tin (for GWC) and specimen cups (for KCl extractions). The person weighing out GWC tins would place finished soils in one pile and tins in another. The labeler would ensure that the GWC finished pile would then be subsampled for KCl extractions into specimen cups as well.
6. NUTRIENT (NH4, NO3, PO4) EXTRACTIONS
Introduction
Exchangeable soil nitrogen (N) pools (NH4 and NO3) are determined by extracting 2 mm sieved soils in 2 M Potassium Chloride (KCl), whereas 0.5 M Sodium Bicarbonate (NaHCO3) is used to extract available phosphorus (PO4).
Materials
● Sieved field moist soil (from gallon bag)
● 2 M KCl (149.1 g KCl to 1 liter DI water)
● 0.5 M NaHCO3 (42 g NaHCO3 to 1 liter DI water)
● Acid washed specimen cups (Fisherbrand Cat#14-828-321)
● Balance
● 50 mL graduated cylinder
● Shaker (Ohaus Cat# 02-106-106)
● 110 mm #1 Whatman filters (WHA1001110)
● 110 mm #40 Whatman filters (WHA1440110)
● Acid washed funnels (Evergreen Scientific Cat# 05-555-6)
● Dixie cups or used specimen cups for pre-leaching filters
● Funnel stand
● Scintillation Vials for extracts (Wheaton Cat# 986701)
NOTE: Be sure to have enough materials for blanks. For each batch of 10 samples, make two blanks (samples with no soil) and process them with the other soil samples.
Procedure for KCl Extractions (day of soil sampling)
1. Place a piece of tape on a clean, acid-washed specimen cup. Label tape with [sample ID]-N and the date.
2. Place labeled specimen cup on scale and tare. Weigh out ~10 g (+/- 0.1 g) of field moist soil into the tared cup. Record exact weight into the spreadsheet.
3. Add 50 mL of 2 M KCl to the cup. Place the lid on top and tightly close.
4. Place closed cups into a 5 gallon bucket. Secure to shaker with wire and shake for one hour. Be sure to include blanks (50 mL KCl in labeled specimen cup, ~20/140 samples) at this step.
5. While samples are shaking, set up funnel stands on the benches. Wearing clean gloves, place an acid washed funnel into the funnel stands for each sample, plus blanks.
6. Fold Whatman 1 filter into each funnel.
7. Place a dixie cup or dirty specimen cup under each funnel. Pre-leach filters two times with 2 M KCl (i.e. fill funnel with KCl and allow to leach into waste cup)
8. Pour leached KCl into the 5 gallon waste container. Wipe down countertops. Place a clean scintillation vial in front of each funnel.
9. Once samples are done shaking, place each sample (soil-KCl slurry) in front of a funnel. Label scintillation vials with sample ID, date, and KCl.
10. Pour off supernatant (liquid part of soil mixture) into the funnel with a filter and collect extract into the labeled scintillation vial. Do not shake the specimen cup before pouring. Only fill the scintillation vial ¾ full to prevent bursting when frozen.
11. Once extracts have been collected, cap scintillation vials and return to the cardboard rack. Label cardboard with “P.I. name, KCl Extracts, Barton, date, plots included, and 1/x racks”
12. Throw away filters. Dump specimen cups into a 5 gallon waste container, then put dirty cups and funnels into plastic tote for cleaning.
13. Wipe down countertops with water first, then 70% EtOH.
14. Store KCl extracts at -20 C until ready to be measured. Transfer sieved soils into fridge (4 C) overnight for PO4 extractions the next morning.
Procedure for PO4 Extractions (morning after soil sampling)
15. On the morning after soil sampling, get bags from the fridge to extract soils for PO4.
16. Repeat the KCl extraction procedure with the following modifications:
a. weigh out 2 g of soil into specimen cups
b. use 40 mL NaHCO3 instead of 50 mL KCl
c. use Whatman 40 filters
d. Label scintillation vials with sample ID, PO4, and date
NOTE: For sampling in Summer 2021, KCl extractions were performed the same day as soil collection following soil sieving. Then samples were placed at 4 C overnight and PO4 was extracted the next morning (within 24 hours of field collection). 140 samples were processed at one time and 15 blanks were added. 2 M KCl and 0.5 M NaHCO3 were made in 10 L batches in 15 L carboys at least a week before extractions to allow complete dissolution. KCl or NaHCO3 were added directly into the carboy with water, and then carboys were shaken vigorously by hand. Solutions sat overnight and then were shaken again until all chemicals were in solution. Prior to sampling day, specimen cups and lids were acid washed and labeling tape was added to the cups.
7. PH & ELECTRICAL CONDUCTIVITY
Introduction
Soil pH is the measure of hydrogen ions in the soil and electrical conductivity (EC) is an indirect measurement of a soil’s salt content.
Materials
● pH/EC meter (Oakton PCTSTestr 50)
● Plastic ‘dixie’ cups
● 400 mL or 1 L waste beaker
● Fresh milliQ water
● Clean graduated cylinder
● Field wet soil (from gallon bags)
● metal spoon
● 70% Ethanol or Isopropyl alcohol to clean spoon between samples
● Parafilm
● Balance
● Kimwipes
Procedure
1. Label plastic ‘dixie’ cup with sample ID. Tare the plastic cup on the balance. Weigh 25 +/- 0.01 g of field wet soil into cup.
2. Using a graduated cylinder, add 25 mL of milliQ water into cup with soil
3. Swirl the cup for one minute to ensure all soil is in solution.
4. Organize all cups in quadrat order on the bench top for easy measuring and recording.
5. After one hour, mix the sample and measure the pH of the soil solution. Be sure the probe is submerged in the solution and hold until the number on the meter stabilizes. Record this number into the spreadsheet. Rinse meter with milliQ squirt bottle in between samples over a waste beaker.
6. After measuring the pH of all samples, swirl the cups to mix again. Switch the pH meter to the EC setting. Place the EC meter into soil, once again ensuring the tip of the probe is submerged and waiting for the reading to stabilize. Rinse meter with milliQ squirt bottle between samples.
7. Once pH and EC measurements are taken/recorded, throw plastic cups away.
NOTE: For sampling in Summer 2021, soil pH and EC were measured the same day PO4 extractions were completed. A smaller team (3-4 personnel) is needed, where 2-3 people weighed soils into plastic cups and other people filled cups with 25 mL of milliQ water from a carboy. Samples were placed in quadrat order on a bench to facilitate easy data recording. Samples were swirled to ensure mixing before sitting for one hour. After samples sat for one hour, samples were mixed again. A team of two would then take pH measurements of each soil (one person measuring, one recording). After pH measurements were taken, EC measurements were taken. Ensure the pH/EC meter is calibrated before use.
8. ARCHIVING SOILS FOR LATER ANALYSIS
Materials
● Sieved soil sample
● Coin envelopes (#6 or #7)
● Scoop or spatula
● KimWipes and isopropyl alcohol
● Sharpie
● Clear lab bench or other space to dry samples
Procedure
After sieved soils have been subsampled for GWC, nutrient extractions, and pH/EC, the remaining soil can be archived for later analysis:
1. Mix the sample well to ensure even distribution of particles.
2. Using a clean spoon to put ~ 40 g of soil (four scoops) into a labeled coin envelope.
3. Clean the spoon with a KimWipe and isopropyl alcohol.
4. Repeat with all samples.
5. Dry at 50 C. Store at room temperature.
NOTE: For sampling in Summer 2021, most of the remaining soil was archived, but a few samples did not have soil remaining to archive. During the procedure, while two people were taking pH/EC measurements, the rest of the team archived the remaining soil (or 40 g) into pre-labeled coin envelopes.
9. IN SITU NITROGEN MINERALIZATION
Introduction
Mineralization is the transformation of organic nitrogen into inorganic nitrogen (NH4 and NO3). At Barton, we measured mineralization by collecting soil cores, placing them in bags, then inserting the bags back into the holes they came from to incubate for a period of time. After this incubation period, we removed the bags and extracted N. By comparing these inorganic N levels to initial inorganic N levels, we can calculate the rate of mineralization.
Materials
● Soil corer
● Pre-labeled 18oz Whirl-pak bags (Cat# B01341)
● Flags
● Trowel
● Gloves
Procedure
Two days after taking time 0 (T0) soil samples (see “Soil Collection” method), additional cores should be collected, placed in Whirl-pak bags, and buried in the ground for 30 days for nutrient extractions:
1. Take two cores 30 cm from the designated edge of each quadrat and place into two pre-labeled Whirl-paks.
2. Ensure there is no plant material in your samples (litter or live plant material). Using a gloved hand, pick out any large plant material.
3. Close each bag and cinch it bag with the cable tie. Make sure the bags are sealed tightly and little to no air is present in the bag.
4. Bury the bags into their respective soil core holes with the end of the cable tie sticking out of the soil surface. Ensure that all of the bag is in the hole, you may need to roll up the bag more.
5. Place a flag near the bag to ease relocating the samples. Be careful not to puncture the bag with the flag.
6. Collect the buried bags after ~28 days of field incubation and bring samples back to the lab on ice for processing.
7. If soils cannot be processed until the next day, leave soil samples in the refrigerator overnight.
8. Discard any bags that have been disturbed by animals or that were punctured when burying. Record into spreadsheet.
9. Sieve soil samples with 2 mm sieve (#10) and transfer the sieved soil into a plastic bag.
10. Label specimen cup and put it on the balance. Tare the balance, weigh 10 g soil, and add 50 mL 2 M KCl. Process the soils using the KCl extraction protocol for available N (NH4/NO3).
11. Label soil tin on tape and weigh. Record into spreadsheet. Add 20-30 g (three spoonfuls) of sieved soil into soil tin. Weigh and record into spreadsheet. Follow gravimetric water content protocol.
NOTE: For sampling in Summer 2021, initial soil cores collected from each quadrat were used as time 0 (T0) (see “Soil Collection” protocol). Mineralization bags were placed 30 cm from the E edge of each quadrat two days after T0 soil collection and left in the field for 49 days before collection.
Mineralization = (ugNO3-N/g dry soil final + ugNH4-N/g dry soil final) - (ugNO3-N/g dry soil initial + ugNH4-N/g dry soil initial)
