Biomass & Necromass > Estimating peak standing stocks of algal biomass and organic matter
Qualitative Analysis of Living and Dead OM in River Reaches
Prepared by Nancy Grimm and Emily Bernhardt
last updated on June 1, 2018
**It would be fabulous to have reach scale estimates of algal biomass and organic matter standing stocks for every portion of your reach over the course of entire years. The work involved in collecting such data is impractical for the majority of researchers. StreamPULSE participants recommend the following approach to collecting this data strategically with the goal of providing qualitative rather than quantified estimates for comparison across sites.
1. Measure peak algal biomass at the time of peak annual GPP
2. Measure peak organic matter standing stocks at the time of peak annual ER
**note - it is possible (and likely) that this will be the same time period for some streams.
3. Make coarse estimates of monthly algal growth rates
4. Make coarse estimates of OM decomposition rates
1. Methods for measuring peak algal biomass at the time of peak annual GPP
If you have a year of metabolism estimates or if you know your river's seasonal cycles particularly well, target this sampling to coincide with the time of peak algal biomass and/or peak GPP. Your goal is to be able to rank your river against other rivers for which annual metabolism has been calculated.
You should delineate your sampling reach based on the flow at the time of sampling so that you are capturing the 'metabolism' footprint at the time. The rule of thumb here is Reach Length = 3v / KO2 where v is velocity (m min-1) and KO2 is the rate of O2 exchange between the water column and the atmosphere (min-1). This distance is the length of river required for 95% turnover of dissolved O2.
Field Equipment:
Laboratory Equipment Needed:
Samples for epilithon standing stock and chlorophyll a are collected using a stratified random sampling design based on the number of channel units determined from your reach characterization (SOP here). You will collect epilithon, filamentous algae and macrophyte biomass from 10 cross sections that are distributed throughout your reach in proportion to the relative length in each channel unit. For example if your reach is 20% riffles, 70% runs and 10% pools you will randomly assign your 10 transects so that there are 2 riffle transects, 7 run transects and 1 pool transect. Use a random # generator to determine the distance upstream of your sensor for each of these transects.
At each transect location lay a line across the stream channel. Collect benthic, floating or rooted material from 5 points distributed across the transect at 10, 30, 50, 70, 90% of the total channel width using the following methods.
Epilithon:
The sample collection procedures differ depending on the bed substrate type.
In the laboratory (< 6 hours from sample collection in the field) the total volume of each slurry sample is determined and recorded. Two well-mixed subsamples should be measured out and filtered through a pre-combusted and tared glass fiber filter (47-mm diameter, Whatman GFF).
AFDM - One filter should be then dried at 60 C, weighed to determine dry mass, combusted (500 C) and reweighed to determine ash-free dry mass. To compute standing stocks of epilithon for the entire stream, weight the averages for each habitat type by the relative proportions of each habitat type in the study reach. Calculate all epilithon standing stocks both in terms of AFDM and dry mass per unit area
Chlorophyll a measurement. The second filter should be placed in a labeled aluminum foil packet and frozen until analysis.
Chlorophyll a will be determined by hot ethanol extraction using the method of Sartory and Grobbelaar (Sartory, D. P. and J. E. Grobbelaar. 1984. Extraction of chlorophyll a from freshwater phytoplankton for spectrophotometric analysis. Hydrobiologia 114:177-187). This method has the benefit of extraction without grinding and avoiding toxic methanol or acetone exposure. Keep samples in the dark at all times, at least in low light when working on them. If the sample is kept in the dark, only 1.3% of the chlorophyll degrades with the 5 min. hot extraction and 24-hour storage. Use test tubes that have the tube numbers scribed on the side with a diamond pencil (ethanol eventually extracts tube numbers written in sharpie, diamond pencils are about $15 at Carolina Biological Supplies) for filter extraction. Place the filter in a container with a known volume (10 mL ethanol in screw cap tubes works well) of 95% ethanol. Mark the location of the meniscus on the side of the tube with sharpie, and place a marble or loose cap on top of the tube. Heat the tube at 79oC for 5 minutes, then mix and cool for 24 h in the dark (can be at room temperature, and can be sealed once cooled if screw cap tubes are used). After extraction, use additional 95% ethanol to bring up to mark on side of tube if ethanol has evaporated, then mix. Clear sample by centrifugation, filtration, or settling. Analyze sample with spectrophotometric analysis at 665 and 750 nm using a 1 cm spectrophotometer cuvette (method in Standard Methods with different extinction coefficient for ethanol and conversion for chlorophyll per unit area). If adsorption is over 1.5 absorbance units, dilute sample. Add 0.1 mL of 0.1 N HCl for each 10 mL of extractant after the first reading and let sample sit for 90 s to phaeophytinize all chlorophyll before reading. Amount of acid is important, too much causes precipitates that you don’t want. Calculations are made as follows:
Chlorophyll a (mg/m2) = (28.78(6650-665a)*v/(A*l) (3)
where 6650 = absorption at 665 before acid addition with absorption at 750 nm subtracted out, 665a = absorption at 665 nm after acid addition with absorption at 750 nm subtracted out, v = volume of extractant used (in liters), A = area of benthos sampled in m2 and l = path length of cell in cm (usually 1 cm).
Phaeophytin (mg/m2) = 28.78 [1.72(665a)-6650]*v/(A*l) (4)
The chlorophyll a data will be used for two purposes: (1) to compute dry mass:chlorophyll a and AFDM:chlorophyll a ratios for those compartments where each of these measurements is made (this will provide a measure of the autotrophic component of epilithon), and (2) to estimate AFDM and dry mass of epilithon in the sand or other habitats where we cannot measure these directly using dry mass:chlorophyll a and AFDM:chlorophyll a ratios.
Macro-autotrophs – standing stock measurements:
Field Equipment:
Cylindrical template (or other template for area to be sampled)
Sample bags (e.g., zip-lock plastic bags)
Laboratory Equipment:
Containers for holding material to be dried (e.g., aluminum pans or trays)
Oven for drying material (60 C)
Oven for combusting material (500 C)
We will determine the standing stock of macro-autotrophs (bryophytes, filamentous algae, macrophytes) by collecting all biomass that is within a circle of 10cm radius of each of the 5 sampling points in each channel cross section. Collection will be relatively straightforward for any attached plants - any macrophyte, bryophyte or filamentous algae that is attached to the streambed within this circle will be collected. For floating algal mats use scissors or a sharp knife to collect a sample of material of known surface area. The material collected is then dried (60 C), weighed, combusted (500 C) and reweighed to determine dry mass and AFDM per unit stream bottom area for each type of material.
**If the distribution of a particular macro-autotroph is highly heterogeneous, then a point/transect survey method may be preferable for determining standing stock. Set up about 20 lateral transects across the stream along the study reach and determine presence or absence at about 10-20 evenly spaced points across each transect. Then sample all material from about 5-10 locations of 100% coverage and measure the area from which the sample was collected. Process the material for dry mass and AFDM as described above, and calculate standing stock per unit stream bottom by multiplying the standing stock in areas of 100% coverage by the % coverage over the entire stream reach.
2. Methods for measuring peak organic matter standing stock at the time of peak ER
We will use a similar approach to estimate OM standing stocks in each stream reach. In cases where this sampling is cooccurring with the autotroph sampling, we recommend putting a transect immediately upstream or downstream of each autotroph sampling reach (as long as you stay within the same channel unit).
Once you have selected your 10 transects use the following approaches to collect particulate organic matter on the surface
Collect surface CPOM: Select points 10, 30, 50, 70 and 90% of the channel widths and collect all of the coarse particulate organic matter (CPOM = leaves, wood) by hand within a 1/2 meter square centered on that point. For very small channels, you may want to simply collect all surface OM within a 1/2 m of your channel cross section tape.
Collect benthic fine particulate organic matter (FBOM) : At each of your sampling points - insert a 5cm diameter 20cm length soil coring tube to the maximum possible depth in the sediments. Cap the core and carefully remove from the sediments. cap the bottom to hold the sediment in place. Once you have up to five cores from your transect - record the depth of sediment in each core
Surficial FBOM: Uncap each core. Use a turkey baster to reduce overlying water to a standard depth over each core (record this water depth). Then use the baster to mix this water and suspend fines on the sediment surface. Use a pipet to collect a well mixed 10mL subsample of overlying water from each core. Combine samples from each core into a single 60mL sample bottle labeled with the transect # and Surface FBOM.
Benthic FBOM: After collecting any surficial FBOM, pour off overlying water from each core. Dump all of the sediments from the transects cores into a bucket and add a known volume of streamwater. Swirl the sediments in the bucket to suspend FBOM. Allow large particles to settle for 10s and then collect a 60mL subsample of the overlying water labeled with the transect # and Benthic FBOM.
** The difficulty of this method will substantially increase with channel depth - as we attempt this in larger systems we are likely to update these methods to allow for traditional boat deployed corers to be used**
In the laboratory filter a known volume of each well mixed FBOM sample through a pre-combusted glass fiber filter (Whatman GFF, 24 mm diameter) making sure there is a thick sediment “cake” on filter. Place filter in a weighed and labeled aluminum weighing pan. Remove CPOM from bags and place in paper bags to dry. Oven dry both the FBOM and CPOM samples in a 60 degree C oven. Weigh CPOM and FPOM samples. After weighing FPOM, combust samples in a muffle furnace at 500 degrees C and reweigh in order to calculate AFDM.
Process samples in order of increasing 15N level (background first, then most downstream station working upstream) in order to minimize possibility for cross contamination of samples. Also process the FBOM material prior to processing the epilithon since the latter is likely to have higher 15N levels. Finally, use a separate filtration device for the upstream samples to avoid potential 15N contamination of these samples. Dry filters (60 C) and set aside in tightly capped scintillation vials (labeled appropriately). Note that we will be getting 15N values per unit dry mass of material sent for analysis (and %N content of dry mass). Therefore, in order to calculate 15N uptake rates, we need measurements of the standing stock of each organic matter compartment in terms of dry mass per unit area of stream bottom (see section 4D).
3. Make coarse estimates of monthly algal growth rates .
We will measure the accumulation of biomass (as AFDM) and chlorophyll on a common substrate (unglazed ceramic tiles) across all sites.
Create three artificial substrate growth platforms for each river reach.
Use aquarium sealant to glue a sheet of unglazed 1" x 1" ceramic tiles (link) onto a plexiglass sheet into which you have drilled two holes.
Install this sheet with the tiles facing up on top of a cinderblock on the streambed. Anchor the platform using rebar or cables attached to the holes in your plexiglass sheet. Try to install at a location that is least likely to dry.
Collect sample tiles from each platform during subsequent visits in order to estimate algal biomass at least four times over the course of a year. At least one sample set should be collected 28 days after deployment (to provide a common measure of algal accrual across all streams).
At each collection date, randomly select 4 tiles from each platform. Pry off. Put two tiles from each platform into each of three foil wrapped and labeled centrifuge tube for subsequent chlorophyll a extraction (as described above for epilithon). Place on ice in the field and store in a freezer until chl a analysis is performed.
Use a toothbrush and a wash bottle to collect attached materials from the other 6 tiles. Put this 'scrubbate' into a single sealable sampling container and cover with Lugol's solution** to allow subsequent algal identification.
**Lugol's solution is commonly used for short-term (e.g. a few months, but possibly a year or more) storage of microalgae. Dissolve one gram of iodine crystals and two grams of potassium iodide in 300 ml of water. Use three drops of this solution in a 100 ml sample (it should look like very weak tea).
4. Make coarse estimates of OM decomposition rates
Wooden ice cream spoons will be deployed in each experimental reach and collected to measure wood decomposition.
Make and installing packets of wooden spoons
Deployment
Sampling Schedule
Processing
Oven dry all spoons overnight at 65 degrees C
Record weights
Use the decline in average weight over time to estimate wood decomposition rate for your reach
Prepared by Nancy Grimm and Emily Bernhardt
last updated on June 1, 2018
**It would be fabulous to have reach scale estimates of algal biomass and organic matter standing stocks for every portion of your reach over the course of entire years. The work involved in collecting such data is impractical for the majority of researchers. StreamPULSE participants recommend the following approach to collecting this data strategically with the goal of providing qualitative rather than quantified estimates for comparison across sites.
1. Measure peak algal biomass at the time of peak annual GPP
2. Measure peak organic matter standing stocks at the time of peak annual ER
**note - it is possible (and likely) that this will be the same time period for some streams.
3. Make coarse estimates of monthly algal growth rates
4. Make coarse estimates of OM decomposition rates
1. Methods for measuring peak algal biomass at the time of peak annual GPP
If you have a year of metabolism estimates or if you know your river's seasonal cycles particularly well, target this sampling to coincide with the time of peak algal biomass and/or peak GPP. Your goal is to be able to rank your river against other rivers for which annual metabolism has been calculated.
You should delineate your sampling reach based on the flow at the time of sampling so that you are capturing the 'metabolism' footprint at the time. The rule of thumb here is Reach Length = 3v / KO2 where v is velocity (m min-1) and KO2 is the rate of O2 exchange between the water column and the atmosphere (min-1). This distance is the length of river required for 95% turnover of dissolved O2.
Field Equipment:
- Wire brush for scraping
- Container with wide mouth to collect epilithon slurry
- Squeeze bottle filled with stream water to wash scrapings and slurry into container
- Paper to trace rock planar areas (or bring rocks back and do planar tracings in lab)
- Sample Bottles/cups to hold individual epilithon samples (either a 100-mL specimen cup or larger bottle, depending on volume of slurry - see text below)
- Plastic bags or bottles for core samples from fine-grained sediment habitats
- Sample template for sampling on bedrock (approx. 2-inch or 4-inch diameter PVC connector with neoprene ring attached to bottom end with silicone sealant)
Laboratory Equipment Needed:
- Pre-combusted glass-fiber filters (Whatman GF/F, 47 mm diameter)
- Filtration set-ups for 47 mm filters
- Hot ethanol (95%) for extraction of chlorophyll
- Spectrophotometer for measurement of chlorophyll
- Oven for drying material (60oC)
- Oven for combusting material (500oC)
Samples for epilithon standing stock and chlorophyll a are collected using a stratified random sampling design based on the number of channel units determined from your reach characterization (SOP here). You will collect epilithon, filamentous algae and macrophyte biomass from 10 cross sections that are distributed throughout your reach in proportion to the relative length in each channel unit. For example if your reach is 20% riffles, 70% runs and 10% pools you will randomly assign your 10 transects so that there are 2 riffle transects, 7 run transects and 1 pool transect. Use a random # generator to determine the distance upstream of your sensor for each of these transects.
At each transect location lay a line across the stream channel. Collect benthic, floating or rooted material from 5 points distributed across the transect at 10, 30, 50, 70, 90% of the total channel width using the following methods.
Epilithon:
The sample collection procedures differ depending on the bed substrate type.
- For bedrock, use a small cylinder of known area (e.g., a short piece of 2-inch PVC pipe) with a foam gasket attached to one end as the sampling template. Push this template firmly against bedrock, vigorously scrape material using a wire brush, and suction scraped material into a bottle (using a turkey baster or large plastic syringe with tip cut off).
- For rocks that can be picked up (e.g., cobble), choose 1-5 rocks (equivalent to a total planar area of about 100-200 cm2) and brush the rock surface thoroughly with a wire brush to loosen the epilithon and slurry the loosened material into a container. After the epilithon has been removed, measure the approximate planar area of each rock brushed by tracing on a piece of paper (use paper from the same batch that has the same weight per unit area), cutting the tracings and weighing them. If you are brushing more than 1 rock to obtain the desired sample area, combine all rocks collected at a particular sampling location into a single “pooled slurry” sample.
- For fine-grained sediments the epilithon will be sampled as FBOM as described in the previous section (4D), unless it exists as a mat in which case it will be included as filamentous algae (see section 4F).
In the laboratory (< 6 hours from sample collection in the field) the total volume of each slurry sample is determined and recorded. Two well-mixed subsamples should be measured out and filtered through a pre-combusted and tared glass fiber filter (47-mm diameter, Whatman GFF).
AFDM - One filter should be then dried at 60 C, weighed to determine dry mass, combusted (500 C) and reweighed to determine ash-free dry mass. To compute standing stocks of epilithon for the entire stream, weight the averages for each habitat type by the relative proportions of each habitat type in the study reach. Calculate all epilithon standing stocks both in terms of AFDM and dry mass per unit area
Chlorophyll a measurement. The second filter should be placed in a labeled aluminum foil packet and frozen until analysis.
Chlorophyll a will be determined by hot ethanol extraction using the method of Sartory and Grobbelaar (Sartory, D. P. and J. E. Grobbelaar. 1984. Extraction of chlorophyll a from freshwater phytoplankton for spectrophotometric analysis. Hydrobiologia 114:177-187). This method has the benefit of extraction without grinding and avoiding toxic methanol or acetone exposure. Keep samples in the dark at all times, at least in low light when working on them. If the sample is kept in the dark, only 1.3% of the chlorophyll degrades with the 5 min. hot extraction and 24-hour storage. Use test tubes that have the tube numbers scribed on the side with a diamond pencil (ethanol eventually extracts tube numbers written in sharpie, diamond pencils are about $15 at Carolina Biological Supplies) for filter extraction. Place the filter in a container with a known volume (10 mL ethanol in screw cap tubes works well) of 95% ethanol. Mark the location of the meniscus on the side of the tube with sharpie, and place a marble or loose cap on top of the tube. Heat the tube at 79oC for 5 minutes, then mix and cool for 24 h in the dark (can be at room temperature, and can be sealed once cooled if screw cap tubes are used). After extraction, use additional 95% ethanol to bring up to mark on side of tube if ethanol has evaporated, then mix. Clear sample by centrifugation, filtration, or settling. Analyze sample with spectrophotometric analysis at 665 and 750 nm using a 1 cm spectrophotometer cuvette (method in Standard Methods with different extinction coefficient for ethanol and conversion for chlorophyll per unit area). If adsorption is over 1.5 absorbance units, dilute sample. Add 0.1 mL of 0.1 N HCl for each 10 mL of extractant after the first reading and let sample sit for 90 s to phaeophytinize all chlorophyll before reading. Amount of acid is important, too much causes precipitates that you don’t want. Calculations are made as follows:
Chlorophyll a (mg/m2) = (28.78(6650-665a)*v/(A*l) (3)
where 6650 = absorption at 665 before acid addition with absorption at 750 nm subtracted out, 665a = absorption at 665 nm after acid addition with absorption at 750 nm subtracted out, v = volume of extractant used (in liters), A = area of benthos sampled in m2 and l = path length of cell in cm (usually 1 cm).
Phaeophytin (mg/m2) = 28.78 [1.72(665a)-6650]*v/(A*l) (4)
The chlorophyll a data will be used for two purposes: (1) to compute dry mass:chlorophyll a and AFDM:chlorophyll a ratios for those compartments where each of these measurements is made (this will provide a measure of the autotrophic component of epilithon), and (2) to estimate AFDM and dry mass of epilithon in the sand or other habitats where we cannot measure these directly using dry mass:chlorophyll a and AFDM:chlorophyll a ratios.
Macro-autotrophs – standing stock measurements:
Field Equipment:
Cylindrical template (or other template for area to be sampled)
Sample bags (e.g., zip-lock plastic bags)
Laboratory Equipment:
Containers for holding material to be dried (e.g., aluminum pans or trays)
Oven for drying material (60 C)
Oven for combusting material (500 C)
We will determine the standing stock of macro-autotrophs (bryophytes, filamentous algae, macrophytes) by collecting all biomass that is within a circle of 10cm radius of each of the 5 sampling points in each channel cross section. Collection will be relatively straightforward for any attached plants - any macrophyte, bryophyte or filamentous algae that is attached to the streambed within this circle will be collected. For floating algal mats use scissors or a sharp knife to collect a sample of material of known surface area. The material collected is then dried (60 C), weighed, combusted (500 C) and reweighed to determine dry mass and AFDM per unit stream bottom area for each type of material.
**If the distribution of a particular macro-autotroph is highly heterogeneous, then a point/transect survey method may be preferable for determining standing stock. Set up about 20 lateral transects across the stream along the study reach and determine presence or absence at about 10-20 evenly spaced points across each transect. Then sample all material from about 5-10 locations of 100% coverage and measure the area from which the sample was collected. Process the material for dry mass and AFDM as described above, and calculate standing stock per unit stream bottom by multiplying the standing stock in areas of 100% coverage by the % coverage over the entire stream reach.
2. Methods for measuring peak organic matter standing stock at the time of peak ER
We will use a similar approach to estimate OM standing stocks in each stream reach. In cases where this sampling is cooccurring with the autotroph sampling, we recommend putting a transect immediately upstream or downstream of each autotroph sampling reach (as long as you stay within the same channel unit).
Once you have selected your 10 transects use the following approaches to collect particulate organic matter on the surface
Collect surface CPOM: Select points 10, 30, 50, 70 and 90% of the channel widths and collect all of the coarse particulate organic matter (CPOM = leaves, wood) by hand within a 1/2 meter square centered on that point. For very small channels, you may want to simply collect all surface OM within a 1/2 m of your channel cross section tape.
Collect benthic fine particulate organic matter (FBOM) : At each of your sampling points - insert a 5cm diameter 20cm length soil coring tube to the maximum possible depth in the sediments. Cap the core and carefully remove from the sediments. cap the bottom to hold the sediment in place. Once you have up to five cores from your transect - record the depth of sediment in each core
Surficial FBOM: Uncap each core. Use a turkey baster to reduce overlying water to a standard depth over each core (record this water depth). Then use the baster to mix this water and suspend fines on the sediment surface. Use a pipet to collect a well mixed 10mL subsample of overlying water from each core. Combine samples from each core into a single 60mL sample bottle labeled with the transect # and Surface FBOM.
Benthic FBOM: After collecting any surficial FBOM, pour off overlying water from each core. Dump all of the sediments from the transects cores into a bucket and add a known volume of streamwater. Swirl the sediments in the bucket to suspend FBOM. Allow large particles to settle for 10s and then collect a 60mL subsample of the overlying water labeled with the transect # and Benthic FBOM.
** The difficulty of this method will substantially increase with channel depth - as we attempt this in larger systems we are likely to update these methods to allow for traditional boat deployed corers to be used**
In the laboratory filter a known volume of each well mixed FBOM sample through a pre-combusted glass fiber filter (Whatman GFF, 24 mm diameter) making sure there is a thick sediment “cake” on filter. Place filter in a weighed and labeled aluminum weighing pan. Remove CPOM from bags and place in paper bags to dry. Oven dry both the FBOM and CPOM samples in a 60 degree C oven. Weigh CPOM and FPOM samples. After weighing FPOM, combust samples in a muffle furnace at 500 degrees C and reweigh in order to calculate AFDM.
Process samples in order of increasing 15N level (background first, then most downstream station working upstream) in order to minimize possibility for cross contamination of samples. Also process the FBOM material prior to processing the epilithon since the latter is likely to have higher 15N levels. Finally, use a separate filtration device for the upstream samples to avoid potential 15N contamination of these samples. Dry filters (60 C) and set aside in tightly capped scintillation vials (labeled appropriately). Note that we will be getting 15N values per unit dry mass of material sent for analysis (and %N content of dry mass). Therefore, in order to calculate 15N uptake rates, we need measurements of the standing stock of each organic matter compartment in terms of dry mass per unit area of stream bottom (see section 4D).
3. Make coarse estimates of monthly algal growth rates .
We will measure the accumulation of biomass (as AFDM) and chlorophyll on a common substrate (unglazed ceramic tiles) across all sites.
Create three artificial substrate growth platforms for each river reach.
Use aquarium sealant to glue a sheet of unglazed 1" x 1" ceramic tiles (link) onto a plexiglass sheet into which you have drilled two holes.
Install this sheet with the tiles facing up on top of a cinderblock on the streambed. Anchor the platform using rebar or cables attached to the holes in your plexiglass sheet. Try to install at a location that is least likely to dry.
Collect sample tiles from each platform during subsequent visits in order to estimate algal biomass at least four times over the course of a year. At least one sample set should be collected 28 days after deployment (to provide a common measure of algal accrual across all streams).
At each collection date, randomly select 4 tiles from each platform. Pry off. Put two tiles from each platform into each of three foil wrapped and labeled centrifuge tube for subsequent chlorophyll a extraction (as described above for epilithon). Place on ice in the field and store in a freezer until chl a analysis is performed.
Use a toothbrush and a wash bottle to collect attached materials from the other 6 tiles. Put this 'scrubbate' into a single sealable sampling container and cover with Lugol's solution** to allow subsequent algal identification.
**Lugol's solution is commonly used for short-term (e.g. a few months, but possibly a year or more) storage of microalgae. Dissolve one gram of iodine crystals and two grams of potassium iodide in 300 ml of water. Use three drops of this solution in a 100 ml sample (it should look like very weak tea).
4. Make coarse estimates of OM decomposition rates
Wooden ice cream spoons will be deployed in each experimental reach and collected to measure wood decomposition.
Make and installing packets of wooden spoons
- Drill a hole one one side of the ice cream spoon and remove splinters/wood dust.
- Dry spoons in a drying oven overnight at 65 degrees C.
- Each spoon should be weighed and tagged (weights recorded by tag #) with a numbered aluminum tree tag attached via cable tie or cord.
- A set of 40 weighed spoons are then placed into a NITEX screen envelope
Deployment
- To deploy: attach the bag of spoons to an anchor in the streambed at a location that is least likely to dry
Sampling Schedule
- Collect 10 spoons on each of 4 dates over the following year
Processing
Oven dry all spoons overnight at 65 degrees C
Record weights
Use the decline in average weight over time to estimate wood decomposition rate for your reach
