Anhydrobiosis

Anhydrobiosis Soils Polygon Study

Abstract: 

In the Antarctic Dry Valleys, soil polygons are prominent features of the landscape and may be key units for scaling local ecological information to the greater region. We examined polygon soils in each of the 3 basins of Taylor Valley, Antarctica. Our objectives were to characterize variability in soil biogeochemistry and biodiversity at local to regional scales, and to test the influence of soil properties upon invertebrate communities. We found that soil biogeochemical properties and biodiversity vary over multiple spatial scales from fine (greater than 10 m) to broad (less than 10 km) scales. Differences in biogeochemistry were most pronounced at broad scales among the major lake basins of Taylor Valley corresponding to differences in geology and microclimate, while variation in invertebrate biodiversity and abundance occurred at landscape scales of 10-500 m, and within individual soil polygons. Variation in biogeochemistry and invertebrate communities across these scales reflects the influence of physical processes and landscape development over ecosystem structure in the dry valleys. The development of soil polygons influences the spatial patterning of soil properties such as soil organic matter, salinity, moisture, and invertebrate habitat suitability. Nematode abundance and life history data indicate that polygon interiors are more suitable habitats than soils in the troughs at the edges of polygons. These data suggest that physical processes (i.e. polygon development) and biogeochemistry are an important influence on the spatial variability of biotic communities in dry valley soil ecosystems.

Data set ID: 

4005

Core Areas: 

Associated Personnel: 

241
242
243

Short name: 

Poly_anhydro

Methods: 

  During the 1995-1996 season, four contingent polygons were selected as P1, P2, P3 and P4 in a clockwise direction. The middle of the south facing crack was selected as the "0" point and soil samples were taken at 0, 0.2, 0.5, 2, and 5 m along a transect towards the middle of the polygon (in a south to north direction).
       
  Soil samples were taken for organism enumeration and moisture content analysis as follows: Sampling bags were prepared with one sterile 'Whirlpak' bag and clean plastic scoop per sample. Samples were taken from within the 85 cm diameter circular area of each plot. The location of the sampling was recorded each year so that areas were not re-sampled. Using the plastic scoop, soil was collected to 10 cm depth. Very large rocks (greater than 20 mm diameter) were excluded from the sample. The soil was shoveled into the 'Whirlpak' bag until three quarters full (about 1.5 kg soil). The soil was mixed well in the bag, then the bag was closed tightly, expelling as much air as possible. The soil samples were stored in a cooler for transportation. On return to the laboratory (within 8 hours of sampling), the soils were stored at 5degreesC until further processing.
     
  In the laboratory, soil samples were handled in a laminar flow hood to prevent contamination. The Whirlpak bags of soil were mixed thoroughly prior to opening. Approximately 200cm3 of soil was placed in a pre-weighed 800mL plastic beaker. Rocks greater than 3-4mm in diameter were removed from the sample. A sub-sample of approximately 50g was removed and placed in a pre-weighed aluminum dish, and weighed on a balance accurate to 0.01g. This sample was dried at 105degreesC for 24 hours. The sample was removed, placed in desiccator to cool down, and re-weighed. These data were used to calculate water content of the soil, and to express data as numbers of soil organisms per unit dry weight of soil.
     
  The remaining soil in the plastic beaker was weighed. Cold tap water was added up to 650 mL. The soil suspension was stirred carefully (star stir or figure of 8) for 30 seconds, using a spatula. Immediately the liquid was poured into wet screens - a stack of 40 mesh on top of a 400 mesh. The screens were rinsed gently with ice cold tap water (from a wash bottle) through the top of the stack, keeping the screens at an angle as the water filtered through. The water was kept on ice at all times. The top screen was removed, and the lower screen rinsed top down, never directly on top of the soil, but at the top of the screen and from behind. The water was allowed to cascade down and carry the particles into the bottom wedge of the angled screen. The side of the screen was tapped gently to filter all the water through. The suspension was rinsed from the front and the back, keeping the screen at an angle and not allowing the water to overflow the edge of the screen. The soil particles were backwashed into a 50mL plastic centrifuge tube, tipping the screen into the funnel above the tube and rinsing the funnel gently. The suspension was centrifuged for five minutes at 1744 RPM. The liquid was decanted, leaving a few mL on top of the soil particles. The tube was filled with sucrose solution (454g sucrose per liter of tap water, kept refrigerated) up to 45mL. This was stirred gently with a spatula until the pellet was broken up and suspended. The suspension was centrifuged for one minute at 1744 RPM, decanted into a wet 500 mesh screen, rinsed well with ice cold tap water and backwashed into a centrifuge tube. Samples were refrigerated at 5degreesC until counted.   
 
   Samples were washed into a counting dish and examined under a microscope at x 10 or x20 magnification. Rotifers and tardigrades were identified and counted. Nematodes were identified to species and sex, and counted. Total numbers in each sample were recorded on data sheets. All species of nematode, and all rotifers and tardigrades found in the sample were recorded. Data were entered in to Excel files, printed, and checked for errors.
    
   Determination of anhydrobiosis. In the laboratory, soil samples were handled in a laminar flow hood to prevent contamination. The Whirlpak bags of soil were mixed thoroughly prior to opening. The remaining soil in the plastic beaker was weighed. 100g of soil (less rocks) were measured into a beaker and brought up to 250 mL with a 1.25M sucrose solution. Mixture was star stirred using a spatula for 30 seconds. Immediately the liquid was poured into screens- a stack of 40 mesh on top of a 400 mesh, prewetted with 1.25M sucrose solution. The screens were rinsed gently with 1.25M sucrose solution (from a wash bottle) through the top of the stack, keeping the screens at an angle as the solution filtered through. The solution was kept on ice at all times. The top screen was removed, and the lower screen rinsed top down, never directly on top of the soil, but at the top of the screen and from behind. The sucrose solution was allowed to cascade down and carry the particles into the bottom wedge of the angled screen. The side of the screen was tapped gently to filter all the solution through. The suspension was rinsed from the front and the back, keeping the screen at an angle and not allowing the solution to overflow the edge of the screen. The bottom screen was then rinsed into a 150 mL beaker with 1.25M sucrose, from the front, using a funnel. The sugar and sediment were then slowly pipetted, using an automatic pipetter, from the beaker into a centrifuge tube containing 10 mL of 2M sugar on ice, at an angle to retain the boundary between sugar layers. The beaker was rinsed with 1.25 sucrose and the dregs were pipetted as well. As many centrifuge tubes as was necessary per sample were used. Each tube was evened off with 1.25M sucrose and centifuged for 5 minutes at 1744 RPM. The supernatant was then poured through a prewetted (with 1.25M sucrose solution) 500 mesh screen, stopping before the sediment came out and rinsed well with ice cold 1.25M sucrose and backwashed into a centrifuge tube. Samples were refrigerated at 5degreesC until counted. 
 
   Samples were washed with 1.25M sucrose into a counting dish and examined under a microscope at x10 or x20 magnification. Nematodes were identified as coiled or uncoiled. Total numbers in each sample were recorded on data sheets. Data were entered in to Excel files, printed, and checked for errors.
 
   Extraction of chlorophyll from the soil. All procedures were carried out in the dark or very low irradiance to avoid degradation of the chlorophyll. The soil samples were mixed thoroughly in the vials, and a sample of approximately 5 g was weighed out in to a 50 mL plastic centrifuge tube with a screw-top cap. 10 mL of a 50:50 DMSO/90% acetone solution was added to each sample and they were mixed thoroughly on a bench-top Vortex mixer for about 5 seconds. The vials were placed in a -4C constant temperature room, in the dark, and left for 12-18 hours.
 
   Determination of chlorophyll a concentration. This was determined fluorometrically using a Turner model 111 fluorometer. A calibration using a known concentration of chlorophyll was carried out prior to sample analysis. The machine was blanked using a 50:50 DMSO/90% acetone solution. Each vial was mixed thoroughly, then centrifuged for 5 minutes at about 1800 RPM. A sample of approximately 4mL of the DMSO/acetone solution was taken from the top of the sample with a pipette, being careful not to get any soil particles in the solution. The sample was placed in a cuvette, in to the fluorometer and the fluorescence was recorded. This was done fairly quickly in order to prevent light from breaking down the chlorophyll. This measurement is called Fo, the initial fluorescence. After taking this reading, 0.1 mL of 1N HCl was added directly to the cuvette and the cuvette was gently agitated. After 20 seconds, the fluorescence was re-measured.(During this step, the acid converts the chlorophyll to phaeophytin by releasing a magnesium ion in an acidic environment). This measurement is called Fa, the fluorescence after acidification. The solution was discarded in to a waste container, and the cuvette rinsed 3 times with DMSO/90% acetone solution before proceeding with the next sample. Data were entered in to Excel files, printed, and checked for errors.

Data sources: 

SOILS_POLY_ANHYDRO

Maintenance: 

This file contains data compiled by Ed Khun, Andy Parsons and Jeb Barrett. The final data QA/QC and analysis were performed by Jeb Barrett.

Metadata was ported to DEIMS by Inigo San Gil (2015)

Additional information: 

Before this dataset was migrated to DEIMS (2015), in the original site we had to construct the columns names: Since the attribute table for this dataset is very large, the user must construct the actual column name.  The FIRST letter represents the species, and the following letters represent the life stage/sex/sum type.  The nermatode species codes are:
      
      S: Scottnema lindsayae 
      E: Eudorylaimus spp. 
      P: Plectus spp.  
      
For example, in the attribute table,  "(code)ML" has the description "The total number of living male (species) adult nematodes extracted from the soil sample in number of organisms per kg soil oven dry weight equivalent. In this case, a column name  called "SML" would be  "The total number of living male Scottnema lindsayae adult nematodes..."
 
Preserved nematode samples are stored in the NREL Preserved Nematode Collection in room 226, College of Natural Resources, Colorado State University. Soils are stored in the Environmental Measurements Laboratory at Dartmouth College.

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