Sapelo Research Application Form
Research Application ID: GCE-134-2023 (submitted: 07/26/2023, status: approved)
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Coastal Connections Bioturbation
Investigator Information
On Island Sponsor: GCE SINERR UGAMI GADNR
| Principal Investigator: | Amanda C. Spivak | ||
| Home Institution: | University of Georgia | ||
| Award Information: | GCE-LTER Supplement award 2023 | ||
| Mailing Address: | Marine Science Rm. 164 | Phone Number: | (706) 542-5709 |
| Marine Sciences | E-mail Address: | aspivak@uga.edu | |
| Athens, Georgia 30602-3636 | |||
| Co-investigators: | |||
Briefly describe the project goals and methodology
Objectives
To determine the effect of bioturbating crabs on invertebrate and microbial communities, and biogeochemistry across latitudes.
To determine how latitude affects the energy content and niche size of crabs across latitudes.
Questions
Question 1: Do crab bioturbators influence carbon stocks, benthic animal and microbial communities similarly across latitudes and wetland types?
Question 2: Does the niche and energy content of crabs vary with latitude and wetland type?
General design for field methods
We will sample in four wetlands on the U.S. east coast: Massachusetts (salt marsh, PIE), Virginia (salt marsh, VCR), Georgia (salt marsh, GCE), Florida (mangrove, FCE).
Invertebrate, microbial and biogeochemistry
For invertebrate and microbial communities and biogeochemistry, samples will be taken from plots (25 cm x 25 cm). We will sample 10 plots along a gradient of burrow densities from 0 to high. Target a variety of densities including at a maximum of three 0-density (no burrow) plots. The goal is to get as many different densities as possible. Plots should be in the same vegetation zone, but at least 3 m from each other. In PIE, VCR, and GCE, focus on the mud fiddler crab, Minuca pugnax. In FCE, focus on the dominant fiddler species in the area.
Sample labels
Site: PIE, VCR, GCE, FCE
Plot number: 1, 2, 3, etc
Burrow densities: numerical value
Food-webs
Food-web sampling will consist of collecting crabs and primary producers at each site. See methods below.
Biogeochemistry and microbial field sampling
Day 1.
Goals: ID sites and quadrats; deploy GHG collars; count burrows and grasses; measure grass height.
1. ID sites and record GPS points; ensure quadrat sampling area and area of the collar have same burrow densities
2. Deploy GHG collars so that the hole is even with the ground
3. Count burrows and plant shoots inside collar
4. Measure stem height in collar
5. Take photo
Day 2:
Goals: Measure GHG fluxes, collect porewater, collect benthic chlorophyll and soil cores, collect microbial samples.
1. Gently place GHG cap on collar; make sure not to disturb collar (lateral wiggles, vertical movement)
2. Attach licor, measure dark fluxes 5 minutes after the line goes linear.; record start and end times, CO2 and CH4 slopes, take screen shot; remove GHG cap
3. Collect PW within collar (but not in a burrow). Put the sipper 7 cm into the soil and attach the 3 way valve. Pull 2 ml into syringe and discard. Pull 62 ml of water. Cap syringe immediately and put on ice in cooler.
4. Collect microbial samples within burrows, adjacent to burrows (edge of the burrow surrounding burrow), and 10 cm outside of burrows. Scrape up to top 5 mm of sediment with a spatula/spoon. Within the burrow, go to a maximum depth of 2.5 cm. Sample burrow first, then adjacent to burrow, then far. Label as 'bur' (within burrow), 'adj' (adjacent to burrow), and far (outside of burrow). Place sediment in cryovials and place in liquid nitrogen.
5. Collect Benthic Chlorophyll in collar. Use cut off 5 ml syringe and collect 3, 1 ml cores into 1 glass scintillation vial. Cap and put in cooler immediately. Keep dark. Freeze ASAP.
6. In adjacent plot with the same burrow density, use Russian peat corer to collect 1, 30 cm core. Open core. Place measuring tape alongside the core with 0 at the top (i.e., marsh surface). Take photo. Use knife to section into 0-5 cm, 5-10 cm, 10-20 cm, and 20-30 cm (bring tape measure. The dimensions are critical.). Completely remove all soil from each core sections to pre-labelled bags using spackle knife. Put on ice and keep cold. Record dimensions and sample id's
Day 2 Post sampling
1. Put data sheets in binder. Take photos of sheets after filtering.
2. Charge licor
3. Benthic chl's in freezer. Keep dark.
4. Microbial samples in freezer.
5. Soil samples in fridge
6. Filter PW
a. Using 2 or 3-way valve transfer 8 ml to a 10 ml syringe
b. Put GFF filters into swinnex with forceps
c. Attach swinnex on 10 ml syringe
d. Push 2 ml of sample H2O through GFF and discard into waste beaker / cup
e. Filter 1 ml into Fe2/3 vial (caution: has concentrated HCl). Cap tightly. invert 3 times. record volume and sample ID
f. Put small tube on to outflow of swinnex; push sample water to end of tube and then lower into H2S vial, below the level of the Zn acetate.
g. Filter 4 ml into H2S vial. Cap tightly. Invert 3 times. Fridge. record volume and sample ID
h. Remove swinnex from 10 ml syringe to 60 ml syringe (w/o small tube)
i. Filter 20 ml into NH4 bottle. Freeze asap. record volume and sample ID
j. Filter water into the pH/redox/ salinity vial so that only 18ml remains in the syringe. Cap vial. Read for pH/redox/ salinity using probes and refractometer after collecting CH4. Record data
k. Remove swinnex from syringe and attach hypodermic needle
l. Carefully pierce the blue butyl septa of the CH4 vial and allow vacuum of the vial to pull in sample. Invert 3 times. record volume and sample ID
Infauna/invertebrate field sampling
1. Count burrow and plant stem density
2. Record any other obvious invertebrates (e.g., mussels, snails)
3. If burrows are present randomly chose a burrow in the plot. Measure its diameter.
4. Take a single 6.6 cm core to a depth of 5 cm on top of that core. Place in pre-labeled Whirl-pak bag. Place in cooler.
5. In lab, add 5% formalin to bags, completely covering sample. Let sit in formalin in Whirl-pak bag for at least 2 days before sieving.
6. For sieving, see JLab's sieving procedure. Or ship to VIMS.
Food-web field sampling
1. Collect 30 large male fiddler crabs (in PIE, VCR, GCE, Minuca pugnax only).
2. Collect primary producers (VCR, GCE, FCE only)
a. Spartina alterinflora - collect 5 leaves of Spartina from different plants (5 leaves total). Place in plastic bag.
b. Filamentous algae
c. Particulate Organic Matter (POM)
d. Benthic microalgae
Where will the project be located?
Dean's Creek marsh: 31.389136°, -81.274270°
How will you provide GPS coordinates for study sites?
GPS coordinates are listed in the project location field
What are the expected start and end dates of the project?
Start Date: 08/07/2023 End Date: 02/29/2024
How many people will access the site and at what frequency?
2 people will sample the sites one time in August. Porewater and greenhouse gas fluxes will be repeated in fall and winter (2 people; 2 days each)
Keywords that describe your project
What equipment will be deployed in the field?
6" PVC collars (10 cm below and above the soil, with 1" holes at the soil surface to allow animal and water movement) will be deployed for 2 days during each sampling trip.
Will plants or animals be collected as part of this study?
Yes: Spartina alterniflora (1 x for isotopes), benthic algae (1 cm cores), Uca crabs (30 males, 1 x), and infaunal invertebrates (1 x)
What are the likely impacts of the project on the site?
There will be minor disturbances due to sample collections but the effects are likely to be short term. we will use minimally invasive techniques, collect minimal sample amounts, and use planks on milk crates as board walks.
Will the project design include boardwalks? If not, explain why not.
We will use planks on milk crates as temporary boardwalks for the day of sampling. the planks will be deployed only when sampling is actively occurring
How long will impacts persist after the research is concluded?
< 1 month
Files attached to this application
GCE-134-2023_Documents_Summary_of_Coastal_Connections_Bioturbation_Project.docx (MS Word file, 13.11 kb, submitted 07/31/2023)
