The Jasper Ridge Global Change Experiment

JRGCE protocols

The following list of protocols roughly decribes techniques used in the JRGCE. Some techniques have varied by year.

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Above-ground harvest protocol
Biomass sorting protocol - species sort
Second biomass harvest sort - (used May 2001)
Second biomass harvest sort - (used May 2002)
Species composition
Microbial biomass C, N, P
Soil extractable inorganic N
Net mineralization cores
Resin available N & P
Litter decomposition

Above-ground harvest protocol

1. Slide harvest area into place, avoiding "gear" within the plots if possible, but being careful not to "choose" the placement.
2. Harvest whole individuals of annual grasses and forbs rooted in the harvest area, being careful to clip as close to the soil surface as possible. This will keep attached (a) all tillers of each individual, and (b) the current year's senescent material.
3. Harvest any piece of the following species which falls within the harvest area, regardless of where it is rooted, i.e. like taking a laser beam vertically from the harvest area and collecting any of these species which fall inside: all perennial grasses, Crepis, Vicia sativa, Dipsicus, Convolvulus. Also, to facilitate identification, place the pieces of perennial grass species into separate labeled bags within the biomass bag.
4. Place plants facing the same direction in the bag, being careful not to dislodge seeds when possible.
5. After all living biomass is removed, collect litter from the harvest square and place in another marked "litter" bag within the biomass bag. Be careful to avoid getting dirt in with the litter, as it creates large errors in the weighing process.
6. "Fluff" the area after you are finished.

Biomass sorting protocol - species sort

1. Sort all individuals to species when possible, being careful to keep all tillers and senescent material attached, this is best accomplished by pulling individuals out from the bundle by their stem bases.
2. Count the number of individuals of each species, place the individuals in an envelope labeled with the species name, plot and quad, harvest date, and number of individuals.
3. Write the number of individuals of each species per harvest square in the summary book.
4. If it is not possible to identify the number of individuals because the tillers were not kept intact, note the number of tillers and write "tillers" after the number in both the summary book and on the envelope for that species.
5. If there are pieces smaller than whole tillers of individuals of a species, note "pieces" on the envelope and in the summary book.
6. If there are grasses with no inflorescence, first try to open the stem to find them, then put them in the "unidentified grass" category, count and label as above.
7. If there are pieces of unidentifiable composites, forbs, or perennial grasses in the harvest bag, put them in the unidentified composite, forb, or perennial categories respectively, count and label as above.
8. Sort through the litter bag to find any green plants or senescent material to add to the species sort. Remove any dirt.
9. Place any senescent material not attached to an identifiable individual in a separate envelope marked senescent.
10. Place all envelopes inside a paper bag marked with the plot and quad into the drying oven.

Second biomass harvest sort - (used May 2001)

1. First sort biomass into four categories: All grass except Avena spp. (both perennial and annual), Avena spp, forbs, and bulbs (Sysrinchium bellum). Late in the season most plants will be senescent and brittle, to maintain whole individuals, it is best to pull individuals out from the bottom of the biomass bundle
2. Separate each pile into senescent (brown, pink, yellow) versus green biomass.
3. Sort through the litter bag to find any green plants or senescent material to add to the sort. Remove any dirt.
4. Place the sorted biomass in an envelope labeled with the category (Avena/grass/forb/bulb/litter), stage (senesent or green), plot, quad, and harvest date.
5. Place all envelopes inside a paper bag marked with the plot and quad into the drying oven.

Second biomass harvest sort - (used May 2002)


Species composition

1. Saw horses are placed on the north and south sides of the plot, and a ladder with pad is placed across the plot on the saw horses. Two plant identifiers lie facing each other on the ladder so they can each look down on two quads.
2. A grid is placed over inside portion of the quad, 10 pins are lowered vertically from fixed points in the grid and pushed gently into the soil. The grid is removed.
3. The identifier lying on the ladder locates the five closest individuals to each pin and calls them off to a recorder.
4. One large perennial plant is one individual, and may not be recorded on multiple pins
5. After the individuals for all 10 pins have been identified, the recorder tallies to assure that 50 individuals total were identified, and the quad is surveyed for any species present in the quad, but not caught by the pin drops. These species are noted as present using checkmarks.

Microbial biomass C, N, P

Briefly, microbial contents are determined using the chloroform-fumigation/extraction technique [W.R. Howarth & E.A. Paul, Microbial Biomass. In Methods of Soil Analysis, Part 2. Microbial and Biochemical Properties - SSSA Book Series No. 5 pp753-773 (1994)]. In this method, fresh soil samples are divided into three subsamples. The first "control" subsample is extracted in salt solution immediately to determine the amount of the element of interest that is available in the soil, the second subsample is fumigated with chloroform to lyse microbial cells and release their contents. This fumigated subsample is then extracted in the same way as the control subsample. The difference in the amount of the element between the first and second samples is the portion of that element contained in the microbial biomass. The third sub-sample is used to calculate and correct for soil moisture. This method is written for 128 samples. It may need to be adapted to allow for soil coring on multiple days, in the past, soil coring has been done in two consecutive days, thus staggering the following procedures.

Microbial C & N

Label plastic bags to collect soil cores
Label 256 (2 sets) of specimen cups for the control & fumigated extractions
Label 256 centrifuge tubes to contain final filtrate of the control & fumigated extractions
Label 256 autoanalyzer vials and organize in styrofoam racks, for analysis of inorganic N on the ALPKEM
Label 128 tin weigh boats for soil moisture calculations
Make freezer and fridge space
Label 128 glass beakers (50 mL) with pencil
Fold 256 pieces of Whatman 42 filter paper in quarters
Make 25 L of 0.5 M K2SO4 in DI water (save left-over to make standards & blanks)
Make sheet for recording weights of subsamples

You will also need:
At least one scale set to measure soil weights to 3 decimal places
4 large vacuum dessicators with additional inserts so samples can be stacked
Vacuum with fresh pump oil, and tubing
4 filtering racks and 48 filter funnels
boiling stones
squeeze bottles for K2SO4
spatula for removing stones
additional 50 mL beakers for chloroform

Timeline for a sample:
Day 1: Collect soil, refrigerate samples until use, subsample soils, extract control soils, start fumigation
Day 2: Filter the control subsamples
Day 5: Extract fumigated soils
Day 6: Filter the fumigated subsamples

Detailed procedure

• Take soil cores to 15 cm, keep cores intact as much as possible, and replace each core from an undisturbed core taken from the buffer zone. Rinse corer between each quad.
• Refrigerate soil until used
• Homogenize soil core by just before subsampling, keep cold as long as possible.

For each quad, subsample as follows:
-Weigh 15-20 g wet soil for "control" into a labeled specimen cup
-Weigh 15-20 g wet soil for fumigated subsample into a numbered glass beaker
-Weigh a numbered tin weigh boat, then add 10 g wet soil and record weight again, and place this subsample into an oven to dry. This dried sample will be re-weighed later, the weight of the boat subtracted, and the intial versus final weight of the soil compared to find the soil moisture content of the subsamples
-If samples are very dry, add 5mL DI water with pipette to the fumigated subsamples after weighing

Extract the control sample by adding 75 mL 0.5 M K2SO4, capping tightly, shaking vigorously for a moment, and placing on a shaker table for one hour, then refrigerate for 24 hours.

Fumigate the subsamples, do all of the following steps in the fume hood:
- Remove ethanol from chloroform by mixing 25 g alumina into 100 mL chloroform, mix with stir bar on plate
- Line dessicator with moist paper towels
- Fill dessicator with samples, no more than 40, leaving room for the chloroform
- Place 40 mL chloroform and several boiling chips in the center of the dessicator, in a ring of foil to prevent any spilled chloroform from getting into the samples
- Attach vacuum pump to closed dessiccator and evacuate until the chloroform begins to boil. Keep boiling for 2 minutes without letting it boil over. Replace air in dessicator. Repeat 5 times boiling for one minute. Seal the dessicator without letting new air in.
- Label dessicator with lab tape indicating the date and time of fumigation
- Cover dessicator with dark cloth or thick garbage bag
- Let sit for 5 days
- On the 5th day, remove lid slowly by sliding in a horizontal plane, remove chloroform and paper towels. Place used chloroform in a labeled jar for hazardous waste disposal.
- Replace cover on dessicator and evacuate 5 times for one minute each to remove chloroform from soil, let air in between each evacuation.

Extract the fumigated samples:
• Transfer the fumigated subsamples to labeled specimen cups, add 75 mL 0.5M K2SO4, cap tightly, shake vigorously, and place on the shaker table for one hour. You may wish to do the transferring of soils in the fume hood, as there may be residual chloroform in the soils.
• Let samples stand overnight in the refrigerator

• After placing folded Whatman 42 filter paper in funnels, rinse the paper with 0.5 M K2SO4
• Pour extracts into funnels and allow them to drip into fresh specimen cups (no need to label these)
• Once all the filtrate has come through, pour some filtrate into a labeled autoanalyzer vial, pour the rest into a labeled 60mL centrifuge vial, freeze samples until analysis.
• Scoop any dirt and rocks left in the funnel back into the original, dirty specimen cup. (Later, wash this through a 2 mm seive, save the rocks, dry and weigh them to correct for rock weight)
• Also pour 75 mL aliquots of K2SO4 through clean filter paper to analyze later.

Analysis of filtrate:
To measure microbial C, you can run these solutions on a TICTOC analyzer (i.e. the Shimadzu in the Matson lab), no digestion or dilution should be necessary. Or, you may choose to use a "dry-down method", where 5 mL of your extract is allowed to dry in a test tube, then the remaining salts are ground and foiballed for analysis on an elemental analyzer, which can be connected to a mass spectrometer. If you use the dry-down method, you can also analyze nitrogen at the same time. Other options for N analysis are Kjeldahl digestion followed by analysis of the digests on an ALPKEM autoanalyzer (this method only gets the NH4 and the organic N); an alkaline persulfate oxidation digest followed by analysis on an ALPKEM (but this method may drive off ammonium). In any case, 3.5 mL subsamples of all filtrate should be saved in autoanalyzer vials and analyzed without digestion to determine inorganic N content.
Note: save samples of DI water, and at least 2 liters of 0.5 M K2SO4 to make standard solutions.

Microbial P

Method for measuring microbial P in 2002 was also based on the chloroform fumigation/extraction method (P.C. Brookes et al 1982, "Measurement of microbial biomass P in soil," Soil Biology & Biochemistry 14:319-329). Adaptations are as follows:
o Prep is the same as for microbial C & N, except that 30 L of 0.5 M NaCO3 should be made instead of the K2SO4.
o Use 10 g wet soil for the control and fumigated subsamples
o Extract control soils in 100 mL 0.5 M NaCO3, shake for 30 minutes on a shaker table, filter on Whatman 42 filter paper rinsed with 0.5 M NaCO3, freeze filtrate.
oFumigate subsamples for 46 hours, extract and filter the same way as controls.
oPerform rock and soil moisture corrections as with microbial C & N.
oFreeze filtrate until analysis on an ICP, no digestion is necessary
oNote: because the ICP uses and acid wash, it would be preferable in the future to use Bray's solution instead of sodium bicarbonate as the extract solution. If you wish to analyze sodium bicarbonate extracts, one drop concentrated HCl should be added to each 6 mL sample in a borosilicate test tube, and racks of tubes should sit in the refrigerator for at least one week, to allow degassing of carbon dioxide.
oNote: to get good P numbers, every piece of glassware should be acid washed, and care should be taken not to contaminate the samples.

Soil extractable inorganic N

oTake cores to 15 cm in the field, keep cold until extraction, extract as soon as possible
oWeigh a 15 g wet soil subsample, record weight. Weigh a second 10 g wet subsample, and place in an oven for 24 hours, later weigh and use the proportional difference to correct for soil moisture.
oExtract subsample in 75 mL 2 M KCl in a specimen cup. Shake vigorously, place on a shaker table for one hour, filter through Whatman 42 filter paper rinsed with 2 M KCl. Save rocks to correct for rock weight. Freeze filtrate in labeled autoanalyzer vials for analysis on an ALPLEM.
oSave some 2M KCl to use in making standards for the ALPKEM

Net mineralization cores

1) Thin –walled PVC tube, 1 3/8" diameter, 15 cm long, with one sharpened end.
2) Plastic caps (1 3/8" inner diameter) with holes cut in the end. (Protec , 800-890-4595, low density polyethylene caps, cat # A-1-3/8). (2 per tube)
3) Mesh screen squares, 2" x 2", (4 per tube)
4) 4 g resin bags in nylons (2 per tube)
5) Duct tape

1) Mark the tube with a sharpie indicating the top and the bottom.
2) Pound tube into the ground out side of the plot in buffer zone to fill tube with soil.
3) Pull tube with soil out of ground
4) Stretch a mesh square across the top
5) Place a resin bag on top of mesh square
6) Stretch a second piece of square mesh over the top of the resin and cap with plastic cap
7) Press down the resin so the top is completely covered in resin.
8) Tape the cap and mesh to the side of the tube with duct tape
9) Do the same for the bottom
10) Place tube into "net mineralization tube" hole in quadrant, right side up.
11) Take a second core nest to the first core in the buffer zone.
12) Put in plastic bag and process for inorganic N.

Resin available N & P

oAnion Exchange resin: AG-1-X8 (Baker or biorad): Exchange capacity 1.2 meq/ml for anions=1.2 mmol/ml resin
oCation exchange resin: HCR-W2 H+ form, spherical beads (16-40 mesh): Exchange capacity is 2.0 meq cations per ml resin=2.0 mmol/ml resin.
oNylon stockings (Leggs in the box work well)
oCable ties
oEar tags
oWhatman vials for extraction

Resin bag assembly:
Batches of mixed anion and cation exchange resins are prepared by blending the beads on a 1:1 ratio according to exchange capacities. Note that they often have a different exchange capacity when wet versus dry - use the wet exchange capacity to balance the ratio. Resins arrive in a wet form, approximately %40 water by weight. Resin beads should dry before weighing, air drying or drying in an oven at 35 degrees C can take several days. The mixed resins are weighed out and placed in knee-hi nylon stockings. Bags are sealed with a small cable tie, and individually labeled using metal "ear tags".

Some cation exchange resins are in the H+ form (check label). When extracted with KCl, the resulting solutions will be highly acidic and can be detrimental to analysis on an autoanalyzer. Therefore, before using the bags it is necessary to remove the H+ by loading them with Na+. To do this, soak in 5 M NaCl overnight. The soaking end point has been reached when no more sodium is taken up, and no more H+ is released into salt solution. Check this by monitoring pH of salt baths after addition of resin bags.

All completed resin bags should be soaked/rinsed with de-ionized water until the conductivity of the rinse water approaches that of tap water. Spin resin bags in a salad spinner to remove excess moisture - 5 turns is sufficient. Store moist resin bags in plastic bags in a refrigerator until placed in the field.

Nutrient Extraction Procedure
1) Collect bags from field site
2) Rinse bags individually w/ de-ionized water to remove soil
3) Spin bags dry individually in salad spinner to remove excess water.
4) Placed resin bag and 50mls of 2MKCl in small plastic or glass bottle with lid. (Volume will vary with size of resin bag: 50mls for 12g bags, 30mls for 5g bags 10mls for 1.5g bags, etc.)
5) Place bottles on shaker for 30 minutes
6) Place two subsamples in labeled vials for analysis on autoanalyzer.
7) Place samples in freezer.

Bag Recycling
After bags are extracted in 2M KCl, they can be placed in several solutions of 5M NaCl to reload the sites with Na+ and Cl- and rinsed with de-ionized water until all the excess salt is removed. Then bags are spun to remove excess moisture and stored in plastic bags until ready to go back to the field

Test for Extraction Efficiency
For any given resin source, bags should be tested to check for possible contamination and for extraction efficiency.
1) From a stock solution with known ppm's of NH4-N and NO3-N, make up a series of loading solutions in a ppm range similar to that anticipated for your samples.
2) After loading, extract bags with 2M KCl. If volume of extracting solution is the same as volume of loading solution, and extraction efficiency is 100%, ppm's loaded will equal ppm's extracted. Actual extraction efficiencies will be from 50-80%.

Litter decomposition

2mm fiberglass mesh coated with nylon (windowscreen, available at any hardware store)
glue guns and glue sticks
ear tags
coin envelopes labeled with: plot, quad, date, and "initial litter for decomposition pucks"

•Harvest senesced litter in late summer by laying a string across quadrants, and taking grass stems that intersect this transect. At least one gram of material should be collected from each quad. Be careful not to break up the litter.
•Dry this litter at 50 degrees C for 24 hours. Note: LTER protocols call for air dried litter to be put in pucks. Future studies should use air dried litter instead.
•Make circular pucks with the mesh and glue to fit in the soil respiration rings, 10 cm in diameter on the bottom, 3 cm high sides
•Label each puck with an ear tag that will match the identity of the litter, and the final location of the puck
•Weigh out a portion of the collected litter proportional to the peak biomass for that quad in the previous year, being careful not to breakup the litter. Record this weight.
•Save the rest of the litter in the labeled coin envelope, later grind it on a Wiley mill (40 mesh), and subsequently powder the sample using a ball grinder, and analyze the powder for initial litter chemistry.
•Place the litter in the puck, being careful not to break the litter or lose any small pieces, and being careful to match the identity of the litter with the label on the puck, and seal the top of the puck with glue.
•Place gently inside the soil respiration ring before the first rains in the fall.
•Collect the puck one year later on a very dry summer afternoon, and carefully place the puck in a labeled plastic bag for transfer to the lab, once again, avoid losing any small peices.
•Cut open the pucks and weigh all remaining litter, being careful to remove any dirt or seeds which got inside. The litter should be dry at this stage.
•Grind this partially decomposed litter in the same manner as the initial litter subsample, and analyze for final litter chemsitry.