US Long-Term Ecological Research Network

LTREB Lake Mývatn Midge Infall 2008-2011

Abstract
Adjacent ecosystems are influenced by organisms that move across boundaries, such as insects with aquatic larval stages and terrestrial adult stages that transport energy and nutrients from water to land. However, the ecosystem-level effect of aquatic insects on land has generally been ignored, perhaps because the organisms themselves are individually small. At the naturally productive Lake Mývatn, Iceland we measured relative midge density on land using passive aerial infall traps during the summers 2008-2011. These traps, a cup with a small amount of lethal preservative, were placed along transects perpendicular to the lake edge and extending ~150-500 m into the shoreline ecosystem and were sampled approximately weekly from May-August. The measurements of midge relative abundance over land were then used to develop a local maximum decay function model to predict proportional midge deposition with distance from the lake (Dreyer et al. <em>in press</em>). In general, peak midge deposition occurrs 20-25 m inland and 70% of midges are deposited within 100 m of shore.
Additional Information
<p>Portions of Abstract and methods edited excerpt from Dreyer et al. <em>in Press</em> which was derived, in part, from these data.</p>
Contact
Dataset ID
306
Date Range
-
Maintenance
On-going
Metadata Provider
Methods
I. Study System Lake Mývatn, Iceland (65&deg;36 N, 17&deg;0&prime; W) is a large (38 km<sup>2</sup>) shallow (4 m max depth) lake divided into two large basins that function mostly as independent hydrologic bodies (Ólafsson 1979). The number of non-biting midge (Diptera: Chironomidae) larvae on the lake bottom is high, but variable: midge production between 1972-74 ranged from 14-100 g ash-free dw m<sup>-2</sup> yr<sup>-1</sup>, averaging 28 g dw m<sup>-2</sup> yr<sup>-1</sup> (Lindegaard and Jónasson 1979). The midge assemblage is mostly comprised of two species (&gt; 90% of total individuals), Chironomus islandicus (Kieffer) and Tanytarsus gracilentus (Holmgren) that feed as larvae in the sediment in silken tubes by scraping diatoms, algae, and detritus off the lake bottom (Lindegaard and Jónasson 1979). At maturity (May-August) midge pupae float to the lake surface, emerge as adults, and fly to land, forming large mating swarms around the lake (Einarsson et al. 2004, Gratton et al. 2008). On land, midges are consumed by terrestrial predators (Dreyer et al. 2012, Gratton et al. 2008), or enter the detrital pool upon death (Gratton et al. 2008, Hoekman et al. 2012). Midge populations naturally cycle with 5-8 year periodicity, with abundances fluctuating by 3-4 orders of magnitude (Einarsson et al. 2002, Ives et al. 2008). II. Midge Infall Measurement We deployed eleven transects of passive, lethal aerial infall traps arrayed at variable distances from Lake Mývatn to estimate relative midge abundance on shore during the summers 2008-2011. Each transect was perpendicular to the lake edge, with traps located at approximately 5, 50, 150, and 500 m (where possible) from shore for a total of 31 traps around the lake. Sampling locations were recorded using GPS and precise distances from the lake were calculated within a geographic information system. Traps consisted of a single 1000 mL clear plastic cup (0.0095 m<sup>2</sup> opening) affixed 1 m above the ground on a stake and filled with 300-500 mL of a 1:1 mixture of water and ethylene glycol and a trace amount of unscented detergent to capture, kill, and preserve insects landing on the surface of the liquid (Gratton et al. 2008, Dreyer et al. 2012). Midges and other insects were emptied from the traps weekly and the traps were reset immediately, thus collections span the entirety of each summer. III. Identification, Counts, and Conversions Midges were counted and identified to morphospecies, small and large. The midge (Diptera,Chrionomidae) assemblage at Mývatn is dominated by two species, Chironomus islandicus (Kieffer)(large, 1.1 mg dw) and Tanytarsus gracilentus (Holmgren)(small, 0.1 mg dw), together comprising 90 percent of total midge abundance (Lindegaard and Jonasson 1979). First, the midges collected in the infall traps were spread out in trays, and counted if there were only a few. Some midges were only identified to the family level of Simuliidae, and other arthropods were counted and categorized as the group, others. Arthropods only identified to the family level Simuliidae or classified as others were not dually counted as Chironomus islandicus or Tanytarsus gracilentus . If there were many midges, generally if there were hundreds to thousands, in an infall trap, subsamples were taken. Subsampling was done using plastic rings that were dropped into the tray. The rings were relatively small compared to the tray, about 2 percent of the area of a tray was represented in a ring. The area inside a ring and the total area of the trays were also measured. Note that different sized rings and trays were used in subsample analysis. These are as follows, trays, small (area of 731 square centimeters), &ldquo;large1&rdquo; (area of 1862.40 square centimeters), and large2 (area of 1247 square centimeters). Rings, standard ring (diameter of 7.30 centimeters, subsample area is 41.85 square centimeters) and small ring (diameter of 6.5 centimeters, subsample area is 33.18 square centimeters). A small ring was only used to subsample trays classified as type &ldquo;large2.&rdquo;The fraction subsampled was then calculated depending on the size of the tray and ring used for the subsample analysis. If the entire tray was counted and no subsampling was done then the fraction subsampled was assigned a value of 1.0. If subsampling was done the fraction subsampled was calculated as the number of subsamples taken multiplied by the fraction of the tray that a subsample ring area covers (number of subsamples multiplied by (ring area divided by tray area)). Note that this is dependent on the tray and ring used for subsample analysis. Finally, the number of midges in an infall trap accounting for subsampling was calculated as the raw count of midges divided by the fraction subsampled (raw count divided by fraction subsampled).Other metrics such as total insects in meters squared per day, and total insect biomass in grams per meter squared day can be calculated with these data. In addition to the estimated average individual midge masses in grams, For 2008 through 2010 average midge masses were calculated as, Tanytarsus equal to .0001104 grams, Chironomus equal to .0010837 grams. For 2011 average midge masses were, Tanytarsus equal to .000182 grams, Chironomus equal to .001268 grams.
Version Number
13

LTREB Lake Mývatn Midge Emergence 2008-2011

Abstract
Adjacent ecosystems are influenced by organisms that move across boundaries, such as insects with aquatic larval stages and terrestrial adult stages that transport energy and nutrients from water to land. However, the ecosystem-level effect of aquatic insects on land has generally been ignored, perhaps because the organisms themselves are individually small. Between 2008-2011 at the naturally productive Lake Myvatn, Iceland we measured total insect emergence from water using emergence traps suspended in the water column. These traps were placed throughout the south basin of Lake Myvatn and were sampled every 1-3 weeks during the summer months (May-August). The goal of this sampling regime was to estimate total midge emergence from Lake Myvatn, with the ultimate goal of predicting, in conjunction with land-based measurements of midge density (see Lake Myvatn Midge Infall 2008-2011) the amount of midges that are deposited on the shoreline of the lake. Estimates from emergence traps between 2008-2011 indicated a range of 0.15 g dw m-2 yr-1 to 3.7 g dw m-2 yr-1, or a whole-lake emergence of 3.1 Mg dw yr-1 to 76 Mg dw yr-1.
Additional Information
<p>Portions of Abstract and methods edited excerpt from Dreyer et al. <em>in Press</em> which was derived, in part, from these data.</p>
Contact
Dataset ID
305
Date Range
-
Maintenance
Ongoing
Metadata Provider
Methods
I. Study System Lake Mývatn, Iceland (65&deg;36 N, 17&deg;0&prime; W) is a large (38 km<sup>2</sup>) shallow (4 m max depth) lake divided into two large basins that function mostly as independent hydrologic bodies (Ólafsson 1979). The number of non-biting midge (Diptera: Chironomidae) larvae on the lake bottom is high, but variable: midge production between 1972-74 ranged from 14-100 g ash-free dw m<sup>-2</sup> yr<sup>-1</sup>, averaging 28 g dw m<sup>-2</sup> yr<sup>-1</sup> (Lindegaard and Jónasson 1979). The midge assemblage is mostly comprised of two species (&gt; 90% of total individuals), Chironomus islandicus (Kieffer) and Tanytarsus gracilentus (Holmgren) that feed as larvae in the sediment in silken tubes by scraping diatoms, algae, and detritus off the lake bottom (Lindegaard and Jónasson 1979). At maturity (May-August) midge pupae float to the lake surface, emerge as adults, and fly to land, forming large mating swarms around the lake (Einarsson et al. 2004, Gratton et al. 2008). On land, midges are consumed by terrestrial predators (Dreyer et al. 2012, Gratton et al. 2008), or enter the detrital pool upon death (Gratton et al. 2008, Hoekman et al. 2012). Midge populations naturally cycle with 5-8 year periodicity, with abundances fluctuating by 3-4 orders of magnitude (Einarsson et al. 2002, Ives et al. 2008). II. Midge Emergence Measurement We used submerged conical traps to estimate midge emergence from Lake Mývatn. Traps were constructed of 2 mm clear polycarbonate plastic (Laird Plastics, Madison, WI) formed into a cone with large-diameter opening of 46 cm (0.17 m<sup>2</sup>). The tops of the cones were open to a diameter of 10 cm, with a clear jar affixed at the apex. The trap was weighted to approximately neutral buoyancy, with the jar at the top containing air to allow mature midges to emerge. Traps were suspended with a nylon line ~1 m below the surface of the lake from an anchored buoy. For sampling, traps were raised to the surface and rapidly inverted, preventing midges from escaping. Jars and traps were thoroughly rinsed with lake water to collect all trapped midges, including unmetamorphosed larvae and pupae, and scrubbed before being returned to the lake to prevent growth of epiphytic algae and colonization by midges. We assume that the emergence traps collect all potentially emerging midges from the sampling area, though it is likely an underestimate, since some midges initially captured could fall out of the trap. Thus, our results should be considered a conservative estimate of potential midge emergence from the surface of the lake.We sampled midge emergence throughout the south basin of Lake Mývatn. Emergence was sampled at six sites in 2008 and 2011 and ten sites in 2009 and 2010, with locations relocated using GPS and natural sightlines. Each site had two traps within 5 m of each other that were monitored during midge activity, approximately from the last week of May to the first week of August. Midge emergence outside of this time frame is extremely low (Lindegaard &amp; Jónasson 1979) and we assume it to be zero. Traps were checked weekly during periods of high emergence (initial and final 2-3 weeks of the study), and bi-weekly during low emergence periods in the middle of the study (July). III. Identification, Counts, and Conversions Midges were counted and identified to morphospecies, small and large. The midge (Diptera,Chrionomidae) assemblage at Mývatn is dominated by two species, Chironomus islandicus (Kieffer)(large, 1.1 mg dw) and Tanytarsus gracilentus (Holmgren)(small, 0.1 mg dw), together comprising 90 percent of total midge abundance (Lindegaard and Jonasson 1979). First, the midges collected in the infall traps were spread out in trays, and counted if there were only a few. Some midges were only identified to the family level of Simuliidae, and other arthropods were counted and categorized as the group, others. Arthropods only identified to the family level Simuliidae or classified as others were not dually counted as Chironomus islandicus or Tanytarsus gracilentus . If there were many midges, generally if there were hundreds to thousands, in an infall trap, subsamples were taken. Subsampling was done using plastic rings that were dropped into the tray. The rings were relatively small compared to the tray, about 2 percent of the area of a tray was represented in a ring. The area inside a ring and the total area of the trays were also measured. Note that different sized rings and trays were used in subsample analysis. These are as follows, trays, small (area of 731 square centimeters), &ldquo;large1&rdquo; (area of 1862.40 square centimeters), and large2 (area of 1247 square centimeters). Rings, standard ring (diameter of 7.30 centimeters, subsample area is 41.85 square centimeters) and small ring (diameter of 6.5 centimeters, subsample area is 33.18 square centimeters). A small ring was only used to subsample trays classified as type &ldquo;large2.&rdquo;The fraction subsampled was then calculated depending on the size of the tray and ring used for the subsample analysis. If the entire tray was counted and no subsampling was done then the fraction subsampled was assigned a value of 1.0. If subsampling was done the fraction subsampled was calculated as the number of subsamples taken multiplied by the fraction of the tray that a subsample ring area covers (number of subsamples multiplied by (ring area divided by tray area)). Note that this is dependent on the tray and ring used for subsample analysis. Finally, the number of midges in an infall trap accounting for subsampling was calculated as the raw count of midges divided by the fraction subsampled (raw count divided by fraction subsampled).Other metrics such as total insects in meters squared per day, and total insect biomass in grams per meter squared day can be calculated with these data. In addition to the estimated average individual midge masses in grams, For 2008 through 2010 average midge masses were calculated as, Tanytarsus equal to .0001104 grams, Chironomus equal to .0010837 grams. For 2011 average midge masses were, Tanytarsus equal to .000182 grams, Chironomus equal to .001268 grams.
Version Number
13

LTREB Biological Limnology at Lake Myvatn 2012-current

Abstract
These data are part of a long-term monitoring program in the central part of Myvatn that represents the dominant habitat, with benthos consisting of diatomaceous ooze. The program was designed to characterize import benthis and pelagic variables across years as midge populations varied in abundance. Starting in 2012 samples were taken at roughly weekly inervals during June, July, and August, which corresponds to the summer generation of the dominant midge,<em>Tanytarsus gracilentus</em>.
Creator
Dataset ID
296
Date Range
-
Maintenance
Ongoing
Metadata Provider
Methods
Benthic Chlorophyll Field sampling (5 samples) (2012, 2013)1. Take 5 cores from the lake2. Cut the first 0.75 cm (1 chip) of the core with the extruder and place in deli container. Label with date and core number.3. Place deli containers into opaque container (cooler) and return to lab. This is the same sample that is used for the organic matter analysis.In 2014, the method for sampling benthic chlorophyll changed. The calculation of chlorophyll was changed to reflect the different area sampled. Below is the pertinent section from the methods protocols. Processing after the collection of the sample was not changed.Take sediment samples from the 5 cores collected for sediment characteristics. Take 4 syringes of sediment with 10mL syringe (15.96mm diameter). Take 4-5cm of sediment. Then, remove bottom 2cm and place top 2cm in the film canister.Filtering1. Measure volume of material in deli container with 60mL syringe and record.2. Homogenize and take 1mL sample with micropipette. The tip on the micropipette should be cut to avoid clogging with diatoms. Place the 1mL sample in a labeled film canister. Freeze sample at negative 20 degrees Celsius unless starting methanol extraction immediately.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec.4. After 6-18 hours, shake container for 5 sec.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 per cent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000 microLiter pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120microLiters of 0.1 N HCl (30microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Chlorophyll Field sampling (5 samples)1. Take 2 samples at each of three depths, 1, 2, and 3m with Arni&rsquo;s zooplankton trap. For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. 2. Empty into bucket by opening the bottom flap with your hand.3. Take bucket to lab.Filtering1. Filter 1L water from integrated water sample (or until the filter is clogged) through the 47 mm GF/F filter. The pressure used during filtering should be low ( less than 5 mm Hg) to prevent cell breakage. Filtering and handling of filters should be performed under dimmed lighting.2. Remove the filter with forceps, fold it in half (pigment side in), and put it in the film canister. Take care to not touch the pigments with the forceps.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec. and place in fridge.4. After 6-18 hours, shake container for 5 sec.5. Analyze sample in fluorometer after 24 hours.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 percent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000uL pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120 microLiters of 0.1 N HCl (30 microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Zooplankton Counts Field samplingUse Arni&rsquo;s zooplankton trap (modified Schindler) to take 2 samples at each of 1, 2, and 3m (6 total). For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. Integrate samples in bucket and bring back to lab for further processing.Sample preparation in lab1. Sieve integrated plankton tows through 63&micro;m mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micro meter mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted as well.6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Benthic Microcrustacean Counts Field samplingLeave benthic zooplankton sampler for 24h. Benthic sampler consists of 10 inverted jars with funnel traps in metal grid with 4 feet. Set up on bench using feet (on side) to get a uniform height of the collection jars (lip of jar = 5cm above frame). Upon collection, pull sampler STRAIGHT up, remove jars, homogenize in bucket and bring back to lab. Move the boat slightly to avoid placing sampler directly over cored sediment.Sample preparation in lab1. Sieve integrated samples through 63 micrometer mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micrometer mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too!6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Chironomid Counts (2012, 2013) For first instar chironomids in top 1.5cm of sediment only (5 samples)1. Use sink hose to sieve sediment through 63 micrometer mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents into small deli container.3. Return label to deli cup (sticking to underside of lid works well).For later instar chironomids in the section 1.5-11.5cm (5 samples)4. Sieve with 125 micrometer mesh in the field.5. Sieve through 125micrometer mesh again in lab to reduce volume of sample.6. Transfer sample to deli container or pitfall counting tray.For all chironomid samples7. Under dissecting scope, pick through sieved contents for midge larvae. You may have to open tubes with forceps in order to check for larvae inside.8. Remove larvae with forceps while counting, and place into a vial containing 70 percent ethanol. Larvae will eventually be sorted into taxonomic groups (see key). You may sort them into taxonomic groups as you pick the larvae, or you can identify the larvae while measuring head capsules if chironomid densities are low (under 50 individuals per taxanomic group).9. For a random sample of up to 50 individuals of each taxonomic group, measure head capsule, see Chironomid size (head capsule width).10. Archive samples from each sampling date together in a single 20mL glass vial with screw cap in 70 percent ethanol and label with sample contents , Chir, sample date, lake ID, station ID, and number of cores. Chironomid Cound (2014) In 2014, the method for sampling chironomid larvae changed starting with the sample on 2014-06-27; the variable &quot;top_bottom&quot; is coded as a 2. In contrast to previous measurements, the top and bottom core samples were combined and then subsampled. Below is the pertinent section of the protocols.Chironomid samples should be counted within 24 hours of collection. This ensures that larvae are as active and easily identified as possible, and also prevents predatory chironomids from consuming other larvae. Samples should be refrigerated upon returning from the field.<strong>For first instar chironomids in top 1.5cm of sediment only (5 samples)</strong>1. Use sink hose to sieve sediment through 63&micro;m mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents using a water bottle into small deli container.3. Return label to deli cup (sticking to underside of lid works well).<strong>For larger instar chironomids in the section 1.5-11.5cm (5 samples)</strong>4. Sieve with 125&micro;m mesh in the field.5. Sieve through 125&micro;m mesh again in lab to reduce volume of sample and break up tubes.6. Transfer sample to deli container with the appropriate label.<strong>Subsample if necessary</strong>If necessary, subsample with the following protocol.a. Combine top and bottom samples from each core (1-5) in midge sample splitter.b. Homogenize sample thoroughly, collect one half in deli container, and label container with core number and &ldquo;1/2&rdquo;c. If necessary, split the half that remains in the sampler into quarters, and collect each in deli containers labeled with core number, &ldquo;1/4&rdquo;, and replicate 1 or 2d. Store all deli containers in fridge until counted, and save until all counting is complete&quot; Chironomid Size (head capsule width) 1. Obtain picked samples preserved in ethanol and empty onto petri dish.2. Sort larvae by family groups, arranging in same orientation for easy measurment.3. Set magnification to 20, diopter, x 50 times4. Take measurments for up to 50 or more individuals of each taxa. Round to nearest optical micrometer unit.5. Fill out data sheet for number of larvae in each taxa, Chironomid measurements for each taxa, date of sample, station sample was taken from, which core the sample came from, who picked the core, and your name as the measurer.6. Enter data into shared sheetSee &quot;Chironomid Counts&quot; for changes in sampling chironomid larvae in 2014.
Version Number
17

LTREB Kalfastrond Peninsula Experiment (KAL) Midge Counts at Lake Myvatn 2008-2011

Abstract
A cross ecosystem resource blocking experiment was conducted on the Kalfastrond peninsula, known as the KAL experiment or KAL midge blocking experiment, at Lake Myvatn to determine the influence of an aquatic resource on a terrestrial food web over time. A manipulative field experiment was used in conjunction with a stable isotope analysis to examine changes in terrestrial arthropod food webs in response to the midge subsidy. Cages were established at 2 by 2 meter plots in 6 blocks spread across the site. Each block included 3 treatment levels, an open control plot, a full exclusion cage and a partial exclusion cage, for a total of 18 experimental plots. Midge exclusion cages were designed to prevent midges from entering plots with such cages. Control open pit midge cages were set as a control which allowed complete access to all arthropods. Partial midge exclusion cages were designed and used to examine any effects of cages themselves on terrestrial responses while minimally affecting midge inputs into the plots and arthropod movement. All cages were set at the middle to end of May to the beginning of August in each year, the period corresponding to the active growing season of plants and the flight activity of midges at this site. Midge activity was measured in all plots to document changes in midge abundance over the course of a season and between years and to assess the degree to which cages excluded midges.Midge abundance in the plots was continuously measured using passive aerial infall traps. Midges from infall traps were counted and identified to morphospecies, where the small species is Tanytarsus gracilentus and the large species is Chironomus islandicus. Some arthropods were only identified to the family level Simuliidae, and other arthropods were lumped in a category named others. If the infall trap contained hundreds to thousands of a particular midge species a subsample for each species was performed to estimate the number of midges trapped. These data are the results of the midge counts from the infall traps.
Contact
Core Areas
Dataset ID
284
Date Range
-
Maintenance
Ongoing
Metadata Provider
Methods
I. Field MethodsThe site where this manipulative field experiment was conducted on the Kalfastrond peninsula at Lake Myvatn is approximately 150 meters long and 75 meters wide. The vegetation consists of grasses Deschampsia spp., Poa spp., and Agrostis spp.), sedges (Carex spp.), and forbs (Ranunculus acris, Geum rivale,and Potentilla palustris). The experimental midge exclusions occurred from the middle or end of May to the beginning of August in each year, the period corresponding to the active growing season of plants and the flight activity of midges at this site. 2 by 2 meter plots were established in 6 blocks spread across the site. Each block included 3 treatment levels, an open control plot, a full exclusion cage and a partial exclusion cage, for a total of 18 experimental plots. Control plots were open to allow complete access to all arthropods. Experimental midge exclusion cages were 1 meter high and constructed from white PVC tubing affixed to rebar posts on each corner of the plot, Plate 1. Full exclusion cages were entirely covered with white polyester netting, 200 holes per square inch, Barre Army Navy Store, Barre VT, USA, to prevent midges from entering the plot. The mesh netting completely enclosed the 2 by 2 by 1 meter frame to prevent flying insects from entering, however the mesh was not secured to the ground in order to allow non flying,ground crawling, arthropods to freely enter and exit the cages. Partial exclusion cages had one 0.5 meter strip of mesh stretched around the outside of the frame and another 0.75 meter strip draped over the top. Partial cages were designed to examine any effects of cages themselves on terrestrial responses while minimally affecting midge inputs into the plots and arthropod movement.The partial exclusion treatment was discontinued in 2011. Each plot contains a pitfall and an infall trap that are continuously sampled during the summer, while the cages are up. Vacuum samples were taken from the plots about once per month in 2008 through 2010 and only once per summer for subsequent summers.Midge activity was measured in all plots to document changes in midge abundance over the course of a season and between years and to assess the degree to which cages excluded midges. Midge abundance in the plots was continuously measured using passive aerial infall traps consisting of a 1000 milliliter clear plastic cup, 95 square centimeter opening, attached to a post 0.5 meters high and filled with 250 milliliters of a 1 to 1 ethylene glycol to water solution and a small amount of unscented detergent to capture and kill insects that alighted upon the surface. Infall traps were emptied about every 10 days.II. AnalysisMidges were counted and identified to morphospecies, small and large. The midge (Diptera,Chrionomidae) assemblage at Myvatn is dominated by two species,Chironomus islandicus (Kieffer)(large, 1.1 mg dw) and Tanytarsus gracilentus(Holmgren)(small, 0.1 mg dw), together comprising 90 percent of total midge abundance (Lindegaard and Jonasson 1979). First, the midges collected in the infall traps were spread out in trays, and counted if there were only a few. Some midges were only identified to the family level of Simuliidae,and other arthropods were counted and categorized as the group, others. Arthropods only identified to the family level Simuliidae or classified as others were not dually counted as Chironomus islandicus or Tanytarsus gracilentus. If there were many midges, generally if there were hundreds to thousands, in an infall trap,subsamples were taken. Subsampling was done using plastic rings that were dropped into the tray. The rings were relatively small compared to the tray, about 2 percent of the area of a tray was represented in a ring. The area inside a ring and the total area of the trays were also measured. Note that different sized rings and trays were used in subsample analysis. These are as follows, Trays, small (area of 731 square centimeters), large1 (area of 1862.40 square centimeters), and large2 (area of 1247 square centimeters). Rings, standard ring (diameter of 7.30 centimeters, subsample area is 41.85 square centimeters) and small ring (diameter of 6.5 centimeters, subsample area is 33.18 square centimeters). A small ring was only used to subsample trays classified as type large2.The fraction subsampled was then calculated depending on the size of the tray and ring used for the subsample analysis. If the entire tray was counted and no subsampling was done then the fraction subsampled was assigned a value of 1.0. If subsampling was done the fraction subsampled was calculated as the number of subsamples taken multiplied by the fraction of the tray that a subsample ring area covers (number of subsamples multiplied by (ring area divided by tray area)). Note that this is dependent on the tray and ring used for subsample analysis. Finally, the number of midges in an infall trap accounting for subsampling was calculated as the raw count of midges divided by the fraction subsampled (raw count divided by fraction subsampled).Other metrics such as total insects in meters squared per day, and total insect biomass in grams per meter squared day can be calculated with these data. in addition to the estimated average individual midge masses in grams, For 2008 through 2010 average midge masses were calculated as, Tanytarsus equal to .0001104 grams, Chironomus equal to .0010837 grams. For 2011 average midge masses were, Tanytarsus equal to .000182 grams, Chironomus equal to .001268 grams.
Version Number
15

North Temperate Lakes LTER: Benthic Macroinvertebrates 1981 - current

Abstract
Macroinvertebrates are collected from selected shoreline and deep water locations in the seven primary lakes (Allequash, Big Muskellunge, Crystal, Sparkling, and Trout lakes, and unnamed lakes 27-02 [Crystal Bog], and 12-15 [Trout Bog]) in the Trout Lake area using modified Hester-Dendy samplers. Samplers are placed at fyke net and gill net locations in August and retrieved 3-4 weeks later. Macroinvertebrates are preserved in ethanol. This dataset contains counts of various groups of macroinvertebrates identified from specific samples. The majority of the identifications are at the genus level. The data table "Benthic Macroinvertebrate Codes" identifies the taxonomic group represented by each group code. Taxonomic references: Ecology and Classification of North American Freshwater Invertebrates, Edited by James H Thorp and Alan P Covich, Academic Press, Inc, 1991; Aquatic Insects of Wisconsin, William L Hilsenhoff, Natural History Museums Council, University of Wisconsin-Madison (1995). Sampling Frequency: annually Number of sites: 7
Core Areas
Dataset ID
11
Date Range
-
Maintenance
Sampling continues, however, sample analysis happens only during specific projects. Samples are maintained in the zoological museum and can be checked out.
Metadata Provider
Methods
Dendies should be placed in the LTER lakes during the last week or immediately following the last week of the LTER fish sampling, and left in the lakes for approximately four weeks. Most of the dendies are set at fyke net sites so that both fish and macroinvertebrate data are collected there, as dendies can provide an indication of the species and biomass available as fish prey. Fish sampling and macrophyte sampling must be finished before dendies are set because their work disturbs the dendy sites. Dendies were set in all the LTER lakes through 1989, and in 1992 and 1993. Only Trout, Crystal and Sparkling lakes were sampled in 1990, 1991, and 1994 to present. Assembly Parts needed for one dendy sampler: Six wide-meshed 3" x 3" vexar mesh squares Four narrow-meshed 3" x 3" vexar mesh squares Two 3" x 3" tempered hardboards One choreboy plastic scrubbing puff One 5 inch long, 1or4" diameter eyebolt with nut Construct the dendy sampler by pushing the eyebolt through the center hole of each piece with the pieces fitting tightly on the bolt so the dendy cannot become compressed. The individual pieces are layered onto the bolt in this order: hardboard, five vexar meshes alternating wide with narrow, choreboy, five vexar meshes alternating wide with narrow, hardboard, nut. Fold the choreboy into three or four layers, with the most frayed end on the inside. The eyebolt should have a zip tie loop through it. Dendies should be assembled at the lab before going out to set them. Placement in Lakes A "dendy set" consists of three dendy samplers from one lake site. In each set, the middle dendy is designated Dendy B and the end dendies as dendies A and C . Dendy sets are placed in the lakes in one of three configurations. Attached set with anchor and subsurface float. Most shoreline sites (also called fyke net sites) are set this way. Each set consists of a 6 meter line with a minnow trap clip at each end and one in the middle. A dendy is attached to each clip, with the middle clip also being attached to a brick anchor with subsurface float. Beginning 2010, the set in Crystal lake is anchored with a dog tieout stake pushed into the sediment. This is an attempt to reduce the migration of Dendies due to camper activity. Unattached set with anchor and subsurface float. Shoreline sites in Sparkling Lakes are set this way. The individual dendies are not attached together. Each dendy has its own anchor, but only the middle dendy has a float. Setting the dendies as unattached sets reduces disturbance and loss of samplers to the curious public on this high use lake. Attached set with surface float and no anchor. Deep hole sites (also called gill net sites) and all bog sites are set this way. Dendies are connected with a 6 meter line as for the shoreline sets, but are not anchored. The three dendies sink at a more equal rate if one of them is not weighted, hopefully coming to rest apart from one another rather than clumped together. Also, an anchor may pull the dendies into the bottom sediments. Shoreline sets are placed parallel to shore in about one meter of water. The floats are 250ml or 500ml plastic bottles partially filled with water so that they remain submerged. Floats should be attached directly to the brick anchors rather than to the dendy or they will pull the dendy off the lake bottom. Floats should be labeled with Trout Lake Station and our phone number. Sets are assembled in the boat and simply tossed over the side at the site. They should be set so that the dendies are spread out the length of the attachment line rather than clumped together. The samplers do not have to be upright. Retrieval Dendy sets remain in the lakes for about four weeks. Select a calm day for retrieval as the dendies and subsurface floats are very hard to see if there are even moderate waves on the lake. Two people make up the retrieval crew: a boat person and a snorkeler. Anchor the boat near the middle dendy, but collect dendies A and C before dendy B. Lying on your belly on the lake bottom, quickly place a dendy in its labeled freezer container, cover with lid, and return it to the boat person. Disturb it as little as possible before getting it contained, and be careful not to drag the other dendies when returning with it to the boat. Due to physical limitations, the deep sites (and bog sites if sampled) are retrieved without snorkeling. Slowly pull the dendies up until they are within reach of the boat person. Place the container in the lake beneath the dendy and lift the dendy from the water with the container. Unclip the line from the dendy, replace the container lid with a mesh panel lid, and drain the lake water from the container. Rinse the mesh lid into the container with 95percent ethanol, filling the container about half full with EtOH. Cover the container, tilting and swirling it to immerse all the invertebrates in ethanol. At each site, note on the field sheet whether all dendies were recovered. Processing Dendies should be processed as soon as possible after collection to avoid desiccation due to evaporation of the ethanol. The freezer container lids do not fit tightly enough to allow long term storage in these containers. Do not allow the samples to freeze; store them in the garage storage room or the gear room if freezing nights are a possibility. Assemble the Dandy Dendy Concentrator (DDC) so the stopper can be removed without crushing any organisms and the drain will flow into the sink. Rinse the DDC thoroughly with tap water. Make sure the stopper is in place, and place a clean beaker or 60 ml sample jar under the insect spout to catch any leaks. Rinse the freezer container lid into the DDC with tap water. Transfer the dendy from the freezer container to a clean two-gallon bucket. The ethanol left in the freezer container can be poured into the DDC now or later. If there is a lot of particulate matter in the ethanol, wait till later to avoid clogging the mesh. Disassemble the dendy over the bucket, rinsing the vexar mesh squares, hardboards, and eyebolt into the bucket under running tap water. Place the mesh, hardboards, and eyebolt into a sample tray or dishpan. Leave the choreboy in the bucket to soak. Using a fine tipped forceps, pick all matter from the mesh pieces, hardboards, and eyebolt, placing it into the DDC. Pick everything even if you don t know what it is, or are sure it s not animal. Rinse all dendy parts again into the bucket, scraping the hardboard with a razor blade under running water. When everything but the choreboy has been picked clean and rinsed, remove the choreboy from the bucket and pour the rinse water through the DDC. Place the bucket below the tap and gently unfold the choreboy. Rinse thoroughly under running tap water, turning the choreboy inside out if the organisms don t rinse clean initially. Pick the choreboy clean with forceps. Pour the rinse water through the DDC and rinse the bucket, sample tray ,and freezer container into the DDC. Wash down the sides of the DDC with tap water. If there is a large amount of material in the DDC, use the forceps to gently transfer some of it to the sample jar. Use 70percent ethanol to rinse the rest of the material into the sample jar. Rinse the DDC stopper and drain spout thoroughly. Label the sample jar in pencil with lake, dendy site number and letter, date set and date retrieved. Also record this information on a lab form and place it in the Dendy 3-ring binder. Fill the sample jar to the neck with 70percent ethanol and cap tightly. Store all dendy samples from the same year together in a cardboard record storage box labeled with Dendy and collection year. Transfer the sample box to the Zoology museum in Madison. Leave all dendy components out to dry, then store in plastic storage boxes. Hardboard pieces need to dry for a long time before storage to ensure they are dry throughout. Store the choreboys in a large plastic bag. Note: To make 4L of 70percent EtOH, mix 2950 ml 95percentEtOH with 1050ml water. Floating The next step in the process would be to separate the plant material from the animal material. This is done by placing the picked material in a pan with a solution of sugar water, in which the animals will float and can be skimmed off the top for identification. Details of this procedure can be found in Anderson, R.O. A Modified Flotation Technique for Sorting Bottom Fauna Samples. Limnology and Oceanography. Vol. 4, pp. 223-225. We do not float LTER samples at this time. They are stored as picked above. &nbsp;</p>
Short Name
NTLIB01
Version Number
35

Landscape Position Project at North Temperate Lakes LTER: Benthic Invertebrate Abundance 1998 - 1999

Abstract
Benthic invertebrate assemblages of 32 lakes were surveyed as part of the Landscape Position Project. We used modified Hester-Dendy colonization substrates to sample benthic invertebrate communities. Each sampling device consisted of a 3&quot;x3&quot; top plate, alternating layers of course and fine mesh, a &#39;&#39;choreboy&#39;&#39; commercial scrubbing puff, alternating layers of coarse (6.35 mm) and fine (3.18 mm) black plastic mesh, and a 3&quot;x3&quot; bottom plate. Two Hester-Dendy samplers were set at a depth of one meter on each of three substrate types (cobble, sand and silt) within each lake for four weeks in late June through late July in either 1998 or 1999. Within each lake, areas of different substrate types were identified using WI-DNR depth contour lake maps, and substrate type was verified by direct observation. Different substrates were sampled to account for invertebrate associations with specific substrate characteristics. Lake order was determined using a modification of the method of Riera et al. (2000). Lake order is a numerical surrogate for groundwater influx and hydrological position along a drainage network, with the highest number indicating the lake lowest in a watershed. Riera, Joan L., John J. Magnuson, Tim K. Kratz, and Katherine E. Webster. 2000. A geomorphic template for the analysis of lake districts applied to Northern Highland Lake District, Wisconsin, U.S.A. Freshwater Biology 43:301-18. Sampling Frequency: one survey on each lake in late June through late July of 1998 or 1999 Number of sites: 32
Core Areas
Dataset ID
96
Date Range
-
Maintenance
completed
Metadata Provider
Methods
We used modified Hester-Dendy colonization substrates to sample benthic invertebrate communities. Each sampling device consisted of a 3&quot;x3&quot; top plate, alternating layers of course and fine mesh, a choreboy commercial scrubbing puff, alternating layers of coarse (6.35 mm) and fine (3.18 mm) black plastic mesh, and a 3&quot;x3&quot; bottom plate. Two Hester-Dendy samplers were set at a depth of one meter on each of three substrate types (cobble, sand and silt) within each lake for four weeks in late June through late July in either 1998 or 1999. Within each lake, areas of different substrate types were identified using WI-DNR depth contour lake maps, and substrate type was verified by direct observation. Different substrates were sampled to account for invertebrate associations with specific substrate characteristics. Lake order was determined using a modification of the method of Riera et al. (2000). Lake order is a numerical surrogate for groundwater influx and hydrological position along a drainage network, with the highest number indicating the lake lowest in a watershed. Riera, Joan L., John J. Magnuson, Tim K. Kratz, and Katherine E. Webster. 2000. A geomorphic template for the analysis of lake districts applied to Northern Highland Lake District, Wisconsin, U.S.A. Freshwater Biology 43:301-18. Sampling Frequency: one survey on each lake in late June through late July of 1998 or 1999 Number of sites: 32
Short Name
LPPINVA1
Version Number
6

Cross Lake Comparison at North Temperate Lakes LTER - Benthic Macroinvertebrates 2003

Abstract
Benthic invertebrates were collected in 2003 as part of Coarse Woody Habitat (CWH) study on 10 Biocomplexity - Cross Lake Comparison lakes in Vilas County, WI.
Core Areas
Dataset ID
231
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
We chose five lakes with high density of houses and five lakes with a low density of houses based on the lakes and housing quintiles developed by Anna Marburg. In each lake, we chose three sites with no wood and three sites with high wood based on the CLC surveys. We took two benthos samples at each site and for the sites with CWH, we took two wood samples. Macroinvertebrates were identified to the lowest possible taxonomic level. Number of sites: 60 Sampling Frequency: once per siteTo characterise the littoral benthic invertebrate community in each lake, invertebrates were collected using an underwater airlift sampler within a 0.25m2 quadrat (see the study by Butkas, Vadeboncoeur &amp; Vander Zanden, 2011, for details about the airlift). This method samples the overall macroinvertebrate community (both epi- and in-faunal species). Samples were collected at a depth of 1 m in triplicate for both sand and cobble habitat in a 500 um mesh bag at the top of the airlift. Samples were transported on ice and hand-sorted within 4 h of collection. Picked specimens were fixed in 70 % ethanol and identified to genus. For statistical analysis, we pooled macroinvertebrate numbers into broad taxonomic groups, namely Tricoptera, Ephemeroptera, Diptera, Amphipoda, Isopoda and Mollusca, because of large among-lake variability in presence at the genus level. Nilsson E, Solomon CT, Wilson KA, Willis TV, Larget B, Vander Zanden MJ. 2012. Effects of an invasive crayfish on benthic invertebrate abundance, fish benthivory and trophic position. Freshwater Biology. 57:10&ndash;23
Short Name
HELMUS2
Version Number
21

Little Rock Lake - CWH Study at North Temperate Lakes LTER - Benthic Macroinvertebrates 2002 - 2004

Abstract
Benthic invertebrates were collected as part of CWH (coarse woody habitat) study on Little Rock Lake in Vilas county, WI. Pre-manipulation sampling of the macroinvertebrate communities was conducted in the summer of 2002 before the CWH reduction and six times after the reduction, in early, mid, and late summer (May-August) of 2003 and 2004. We divided the shoreline of Little Rock Lake into 50 m sections and randomly chose five sections from each basin for each separate sampling of macroinvertebrates. We collected two benthos and two CWH macroinvertebrate samples at each section. We constructed a benthos sampler by connecting a SCUBA tank to a 7.6 cm PVC pipe with a hose attached 10 cm from one end of the pipe (Wahle and Steneck 1991; Roth et al. 2007). A 500 m Nitex mesh bag was place at the top end of the pipe furthest from the attached hose. Once the tank was turned on, a vacuum formed that sucked the benthos sample into the bag. We used a 0.09 m2 hoop to delineate the benthos sampling area. We sampled CWH using a self-contained, battery-powered aquatic vacuum with a 500 m Nitex mesh bag (Vander Zanden et al. 2006). Sampling lasted for 30 seconds. All samples were stored in 95% ethanol until processed. Macroinvertebrates were identified to the lowest possible taxonomic level. Helmus M.R. and Sass G.G. (2008) The rapid effects of a whole-lake reduction of coarse woody debris on fish and benthic macroinvertebrates. Freshwater Biology, 53, 1423-1433 Number of sites: 44 Sampling Frequency: once pre-manipulation, 6 sampling regimes after reduction
Core Areas
Dataset ID
230
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
Benthic invertebrates were collected as part of CWH (coarse woody habitat) study on Little Rock Lake in Vilas county, WI. Pre-manipulation sampling of the macroinvertebrate communities was conducted in the summer of 2002 before the CWH reduction and six times after the reduction, in early, mid, and late summer (May-August) of 2003 and 2004. We divided the shoreline of Little Rock Lake into 50 m sections and randomly chose five sections from each basin for each separate sampling of macroinvertebrates. We collected two benthos and two CWH macroinvertebrate samples at each section. We constructed a benthos sampler by connecting a SCUBA tank to a 7.6 cm PVC pipe with a hose attached 10 cm from one end of the pipe (Wahle and Steneck 1991; Roth et al. 2007). A 500 m Nitex mesh bag was place at the top end of the pipe furthest from the attached hose. Once the tank was turned on, a vacuum formed that sucked the benthos sample into the bag. We used a 0.09 m2 hoop to delineate the benthos sampling area. We sampled CWH using a self-contained, battery-powered aquatic vacuum with a 500 m Nitex mesh bag (Vander Zanden et al. 2006). Sampling lasted for 30 seconds. All samples were stored in 95% ethanol until processed. Macroinvertebrates were identified to the lowest possible taxonomic level. Helmus M.R. and Sass G.G. (2008) The rapid effects of a whole-lake reduction of coarse woody debris on fish and benthic macroinvertebrates. Freshwater Biology, 53, 1423-1433 Number of sites: 44 Sampling Frequency: once pre-manipulation, 6 sampling regimes after reduction
Short Name
HELMUS1
Version Number
25

Food Web Isotope Study at North Temperate Lakes LTER 2004 - 2010

Abstract
Stable isotope ratios (d13C and d15N) were measured in archived scale samples of seven fish species captured in Sparkling Lake over the period 1981 through 2009. Stable isotopes of benthic and pelagic food web end members (macroinvertebrates and zooplankton, respectively) were also measured. Zooplankton were from samples taken in Sparkling Lake, Allequash Lake, and Big Muskellunge Lake. Zoobenthos were from samples taken in Sparkling Lake. Isotope Study abstract paragraph 2 Number of sites: 3
Core Areas
Dataset ID
261
Date Range
-
Maintenance
completed
Metadata Provider
Methods
Stable isotope ratios (d13C and d15N) were measured in archived scale samples of seven fish species captured in Sparkling Lake over the period 1981 through 2009. Stable isotopes of benthic and pelagic food web end members (macroinvertebrates and zooplankton, respectively) were also measured. Zooplankton were from samples taken in Sparkling Lake, Allequash Lake, and Big Muskellunge Lake. Zoobenthos were from samples taken in Sparkling Lake.
Short Name
CTSISO
Version Number
18
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