US Long-Term Ecological Research Network

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

Little Rock Lake Experiment at North Temperate Lakes LTER: Zooplankton count 1983 - 2000

Abstract
The Little Rock Acidification Experiment was a joint project involving the USEPA (Duluth Lab), University of Minnesota-Twin Cities, University of Wisconsin-Superior, University of Wisconsin-Madison, and the Wisconsin Department of Natural Resources. Little Rock Lake is a bi-lobed lake in Vilas County, Wisconsin, USA. In 1983 the lake was divided in half by an impermeable curtain and from 1984-1989 the northern basin of the lake was acidified with sulfuric acid in three two-year stages. The target pHs for 1984-5, 1986-7, and 1988-9 were 5.7, 5.2, and 4.7, respectively. Starting in 1990 the lake was allowed to recover naturally with the curtain still in place. Data were collected through 2000. The main objective was to understand the population, community, and ecosystem responses to whole-lake acidification. Funding for this project was provided by the USEPA and NSF. Zooplankton samples are collected from the treatment and reference basins of Little Rock Lake at at two to nine depths using a 30L Schindler Patalas trap (53um mesh). Zooplankton samples are preserved in buffered formalin and archived. Data are summed over sex and stage and integrated volumetrically over the water column to provide a lake-wide estimate of organisms per liter for each species. Sampling Frequency: varies - Number of sites: 2
Core Areas
Dataset ID
251
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
We collect zooplankton samples at the deepest part of the lake using two different gear types. We take one vertical tow with a Wisconsin Net (80um mesh), and a series of Schindler Patalas (53um mesh) samples spanning the water column. All samples are preserved in cold 95percent EtOH. After collection we combine subsamples of the individual Schindler Patalas trap samples to create one hypsometrically pooled sample for each lakeordate. The individual depth samples are discarded after pooling except from one August sampling date per year. The Hypsometrically Pooled sample and the Wisconsin Net sample are archived in the UW Zoology museum. We count zooplankton in one or two subsamples, each representing 1.8L of lake water, of the hypsometrically pooled samples to calculate zooplankton abundance. We count one sample date per month from the open water season, and the February ice cover sample. We identify individuals to genus or species, take length measurements, and count eggs and embryos. Protocol log: 1981-May1984 -- a 0.5m high, 31L Schindler Patalas trap with 80um mesh net was used. Two Wisconsin Net tows were collected. Preservative was 12percent buffered formalin. June1984 -- changed to 53um mesh net on Schindler trap. July1986 -- began using the 2m high, 45L Schindler Patalas trap. Changed WI Net collection to take only one tow. 2001 -- changed zooplankton preservative from 12percent buffered formalin to 95percent EtOH. The number of sample dates per year counted varies with lake and year, from 5 datesoryear to 17 datesoryear. 1981-1983 -- pooled samples are of several types: Total Pooled (TP) were created using equal volume subsamples of the Schindler samples. Epi, Meta, Hypo pooled used equal volume subsamples from the Schindler samples collected from each of the thermal strata. Strata Pooled used equal volume subsamples from the Epi, Meta, Hypo pooled samples to create an entire lake sample. Hypsometrically Pooled (HP) is our standard, which uses subsample volumes weighted to represent the hypsometry of the lake.
Short Name
LRZOOP1
Version Number
3

Cascade Project at North Temperate Lakes LTER: Phytoplankton 1984 - 1995

Abstract
Data on epilimnetic phytoplankton from 1984-95, determined by light microscopy from pooled Van Dorn samples at 100percent, 50percent, and 25percent of surface irradiance. There have been 4 counters during this period, with the same counter from 1991-95. Standardization among counters is difficult, so I recommend sticking to the 1991-95 data if possible. Cottingham (1996) describes the counting protocols in detail. Sampling Frequency: varies Number of sites: 5
Core Areas
Dataset ID
80
Date Range
-
LTER Keywords
Maintenance
completed
Metadata Provider
Methods
See for detail: Cottingham, K.L., and S.E. Knight. 1995. Effects of grazer size on the response of mesotrophic lakes to experimental enrichment. Water Science and Technology 32(4): 157-163.
Short Name
CPHYT1
Version Number
3

Biocomplexity at North Temperate Lakes LTER; Coordinated Field Studies: Zooplankton Presence/Absence 2001 - 2004

Abstract
Zooplankton samples were taken at approximately the deepest part of 58 lakes included in the &quot;cross-lake comparison&quot; segment of the Biocomplexity Project. The samples were from years 2001 through 2004. The study lakes are located in Vilas County, Wisconsin and were chosen to represent a range of positions on gradients of both human development and landscape position. Zooplankton samples were analyzed for planktonic crustacean and insect species. Number of sites: 58 Sampling Frequency: each site sampled once
Core Areas
Dataset ID
208
Date Range
-
Maintenance
completed
Metadata Provider
Methods
Wisconsin Net samplesLower the Wisconsin net to the bottom sample depth ( top of the net should be one meter above the bottom). Pull it up slowly at a rate of about 3 seconds per meter. A slow haul prevents the net from pushing water and plankton away from the mouth of the net. To drain the cup swirl it until the water level is below the lower mesh window, then pour contents into the sample jar. Avoid inverting the cup while swirling, as you will lose the sample into the net. Rinse the inside of the cup with 95percent ETOH several times adding the rinse to the sample jar. Wait until the chemistry crew member is finished taking Temp or D.O. profile before taking the Wisconsin net sample, so as not to stir up the sediments. Take replicate sample.
Short Name
BIOZOOP1
Version Number
7

North Temperate Lakes LTER: Zooplankton - Madison Lakes Area 1997 - current

Abstract
Zooplankton samples for the 4 southern Wisconsin LTER lakes (Mendota, Monona, Wingra, Fish) have been collected for analysis by LTER since 1995 (1996 Wingra, Fish) when the southern Wisconsin lakes were added to the North Temperate Lakes LTER project. Samples are collected as a vertical tow using an 80-micron mesh conical net with a 30-cm diameter opening (net mouth: net length ratio = 1:3) consistent with sampling conducted by the Wisconsin Dept. Natural Resources in prior years. Zooplankton tows are taken in the deep hole region of each lake at the same time and location as other limnological sampling; zooplankton samples are preserved in 70% ethanol for later processing. Samples are usually collected with standard tow depths on most dates (e.g., 20 meters for Lake Mendota) but not always, so tow depth is recorded as a variate in the database. Crustacean species are identified and counted for Mendota and Monona and body lengths are recorded for a portion of each species identified (see data protocol for counting procedure); samples for Wingra and Fish lakes are archived but not routinely counted. Numerical densities for Mendota and Monona zooplankton samples are reported in the database as number or organisms per square meter without correcting for net efficiency. [Net efficiency varies from a maximum of about 70% under clear water conditions; net efficiency declines when algal blooms are dense (Lathrop, R.C. 1998. Water clarity responses to phosphorus and Daphnia in Lake Mendota. Ph.D. Thesis, University of Wisconsin-Madison.)] Organism densities in number per cubic meter can be obtained by dividing the reported square-meter density by the tow depth, although adjustments for the oxygenated depth zone during the summer and early fall stratified season is required to obtain realistic zooplankton volumetric densities in the lake's surface waters. Biomass densities can be calculated using literature formulas for converting organism body lengths reported in the database to body masses. Sampling Frequency: bi-weekly during ice-free season from late March or early April through early September, then every 4 weeks through late November; sampling is conducted usually once during the winter (depending on ice conditions). Number of sites: 4 Note: for a period between approximately 2011 and 2015, a calculation error caused density values to be significantly greater than they should have been for the entire dataset. That issue has been corrected.
Core Areas
Dataset ID
90
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
We collect zooplankton samples at the deepest part of the lake using two different gear types. We take one vertical tow with a Wisconsin Net (80um mesh), and a series of Schindler Patalas (53um mesh) samples spanning the water column. All samples are preserved in cold 95percent EtOH. After collection we combine subsamples of the individual Schindler Patalas trap samples to create one hypsometrically pooled sample for each lakeordate. The individual depth samples are discarded after pooling except from one August sampling date per year. The Hypsometrically Pooled sample and the Wisconsin Net sample are archived in the UW Zoology museum. We count zooplankton in one or two subsamples, each representing 1.8L of lake water, of the hypsometrically pooled samples to calculate zooplankton abundance. We count one sample date per month from the open water season, and the February ice cover sample. We identify individuals to genus or species, take length measurements, and count eggs and embryos. Protocol log: 1981-May1984 -- a 0.5m high, 31L Schindler Patalas trap with 80um mesh net was used. Two Wisconsin Net tows were collected. Preservative was 12percent buffered formalin. June1984 -- changed to 53um mesh net on Schindler trap. July1986 -- began using the 2m high, 45L Schindler Patalas trap. Changed WI Net collection to take only one tow. 2001 -- changed zooplankton preservative from 12percent buffered formalin to 95percent EtOH. The number of sample dates per year counted varies with lake and year, from 5 datesoryear to 17 datesoryear. 1981-1983 -- pooled samples are of several types: Total Pooled (TP) were created using equal volume subsamples of the Schindler samples. Epi, Meta, Hypo pooled used equal volume subsamples from the Schindler samples collected from each of the thermal strata. Strata Pooled used equal volume subsamples from the Epi, Meta, Hypo pooled samples to create an entire lake sample. Hypsometrically Pooled (HP) is our standard, which uses subsample volumes weighted to represent the hypsometry of the lake.
Short Name
NTLPL06
Version Number
31

North Temperate Lakes LTER: Phytoplankton - Madison Lakes Area 1995 - current

Abstract
Phytoplankton samples for the 4 southern Wisconsin LTER lakes (Mendota, Monona, Wingra, Fish) have been collected for analysis by LTER since 1995 (1996 Wingra, Fish) when the southern Wisconsin lakes were added to the North Temperate Lakes LTER project. Samples are collected as a composite whole-water sample and are preserved in gluteraldehyde. Composite sample depths are 0-8 meters for Lake Mendota (to conform to samples collected and analyzed since 1990 for a UW/DNR food web research study), and 0-2 meters for the other three lakes. A tube sampler is used for the 0-8 m Lake Mendota samples; samples for the other lakes are obtained by collecting water at 1-meter intervals using a Kemmerer water sampler and compositing the samples in a bucket. Samples are taken in the deep hole region of each lake at the same time and location as other limnological sampling. Phytoplankton samples are analyzed by PhycoTech, Inc., a private lab specializing in phytoplankton analyses (see data protocol for procedures). Samples for Wingra and Fish lakes are archived but not routinely counted. Permanent slide mounts (3 per sample) are prepared for all analyzed Mendota and Monona samples as well as 6 samples per year for Wingra and Fish; the slide mounts are archived at the University of Wisconsin - Madison Zoology Museum. Phytoplankton are identified to species using an inverted microscope (Utermohl technique) and are reported as natural unit (i.e., colonies, filaments, or single cells) densities per mL, cell densities per mL, and algal biovolume densities per mL. Multiple entries for the same species on the same date may be due to different variants or vegetative states - (e.g., colonial or attached vs. free cell.) Biovolumes for individual cells of each species are determined during the counting procedure by obtaining cell measurements needed to calculate volumes for geometric solids (e.g., cylinders, spheres, truncated cones) corresponding to actual cell shapes. Biovolume concentrations are then computed by mulitplying the average cell biovolume by the cell densities in the water sample. Note that one million cubicMicrometers of biovolume PerMilliliter of water are equal to a biovolume concentration of one cubicMillimeterPerMilliliter. Assuming a cell density equal to water, a cubicMillimeterPerMilliliter of biovolume converts to a biomass concentration of one milligramPerLiter. Sampling Frequency: bi-weekly during ice-free season from late March or early April through early September, then every 4 weeks through late November; sampling is conducted usually once during the winter (depending on ice conditions). Number of sites: 4Several taxonomic updates have been made to this dataset February 2013, see methods for details.
Dataset ID
88
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
Water samples are taken along routine sampling and then prepared into permanent slides by the company Phyco Tech. Slides are available for all years, however, species may not have been determined for all available slides.several taxonomic updates were implemented in February 2013, this includes simple name changes to currently accepted names, changes from genus level to species based on long term experience by Phyco Tech, and some slides were revisited to resolve taxonomic uncertainty.1) Converted all Melosira entries to Aulacoseira. The species names have been changed appropriately. 2) Converted all Oscillatoria entries to Psuedanabaena. The species names have been changed appropriately. 3) Converted all Synedra tenera to Synedra filiformis. 4) Converted all Phacotus entries without a species name to Phacotus
lendneri. 5) Converted all Phormidium mucicola to Psuedanabaena 6) Converted Glenodinium entries without a species name to
Glenodinium quadridens 7) Assume that all other entries with genera names but not species
names cannot be resolved to species. 8) Converted all Chrysococcus entries to Chrysocccus minutus 9) Changed some single-celled Microcystis entries so that they would match the format of the colonial entries (genus + species) 10) Resolved some entries to species that were previously coded incorrectly by genus. 11) Added in Cylindrospermopsis raciborskii entries that were recently recounted and changed from Anabaenopsis raciborskii. 12) Converted all entries of genus Erkenia to Erkenia subaequiciliata
Short Name
NTLPL05
Version Number
29

North Temperate Lakes LTER: Phytoplankton - Trout Lake Area 1984 - current

Abstract
Phytoplankton samples from the seven northern Wisconsin LTER lakes in the Trout Lake area (Allequash, Big Muskellunge, Crystal, Sparkling, and Trout lakes and bog lakes 27-02 [Crystal Bog], and 12-15 [Trout Bog]) are collected six times per year at the deep hole sampling station at the same time as our other limnological sampling is conducted. We use a peristaltic pump and tubing, collecting a separate sample from the epilimnion, metalimnion and hypolimnion for most of the lakes. For 27-2 Bog Lake, which is only 2m deep, we collect one 0-2m composite sample. The samples are preserved with Lugols iodine solution. We create a single hypsometrically pooled composite sample per lake from subsamples of the strata samples. The pooled samples are sent to PhycoTech, Inc., a private lab specializing in phytoplankton analysis, to be made into permanent slide mounts. The slide mounts, 3 slides per sample, are archived at the University of Wisconsin - Madison Zoology Museum Phytoplankton are identified to species using an inverted microscope (Utermohl technique) and are reported as natural unit (i.e., colonies, filaments, or single cells) densities per mL, cell densities per mL, and algal biovolume densities per mL. Multiple entries for the same species on the same date may be due to different variants or vegetative states - (e.g., colonial or attached vs. free cell.) Biovolumes for individual cells of each species are determined during the counting procedure by obtaining cell measurements needed to calculate volumes for geometric solids (e.g., cylinders, spheres, truncated cones) corresponding to actual cell shapes. Biovolume concentrations are then computed by mulitplying the average cell biovolume by the cell densities in the water sample. Note that one million cubicMicrometers of biovolume PerMilliliter of water are equal to a biovolume concentration of one cubicMillimeterPerMilliliter. Assuming a cell density equal to water, a cubicMillimeterPerMilliliter of biovolume converts to a biomass concentration of one milligramPerLiter. Sampling Frequency: 6 samples per year Number of sites: 7
Dataset ID
238
Date Range
-
LTER Keywords
Maintenance
ongoing
Metadata Provider
Methods
Water samples are taken along routine sampling and then prepared into permanent slides by the company Phyco Tech. Slides are available for all years, however, species may not have been determined for all available slides.
Short Name
NTLPL08
Version Number
19

North Temperate Lakes LTER: Zooplankton - Trout Lake Area 1982 - current

Abstract
Zooplankton samples are collected from the seven primary northern lakes (Allequash, Big Muskellunge, Crystal, Sparkling, and Trout lakes and bog lakes 27-02 [Crystal Bog], and 12-15 [Trout Bog]) at two to nine depths using a 2 m long Schindler Patalas trap (53um mesh) and with vertical tows (1 m above the bottom of the lake to the surface) using a Wisconsin net (80um mesh). Zooplankton samples are preserved in buffered formalin (up until the year 2000) or 80% ethanol (2001 onwards) and archived. Data are summed over sex and stage and integrated volumetrically over the water column to provide a lake-wide estimate of organisms per liter for each species. A minimum of 5 samples per lake-year are identified and counted. Sampling Frequency: fortnightly during ice-free season - every 6 weeks during ice-covered season. Number of sites: 7.
Core Areas
Dataset ID
37
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
Sample Collection: Schindler-Patalas Trap For LTER lakes use the 2-meter high, 45L Schindler-Patalas trap with 53um mesh net and cup. The volume of the trap used should be indicated on the Volume by Weight form. Collect samples from the target depths at the deep sampling station in each lake. Sample depths are measured from the middle of the trap. Target Depths: TR: 1, 3, 5, 7, 9, 15, 20, 27, 31 meters CRorBM: 1, 3, 5, 7, 9, 11, 13, 15, 18 meters SP: 1, 3, 5, 7, 9, 11, 13, 15, 17 meters ALorTB: 1, 3, 6 meters CB: 1 meter Take samples starting at the surface and working down. Lower the trap slowly so that it remains vertical in the water. Pause at the target depth long enough to allow both trap doors to close completely, and check when it reaches the surface that both did close. Drain the trap through the net and cup, swirling the cup until the liquid level is below the mesh windows. Remove the cup from the net and pull out the center pin to drain the sample into jar, then rinse cup and pin several times with 95percent EtOH into the sample jar. Sample Collection: Wisconsin Net Lower the net to the bottom sample depth. Pull it up slowly, at a rate of about 3 seconds per meter. A slow haul prevents the net from pushing water and plankton away from the mouth of the net. Drain the cup until the water level is below the lower window, then pour contents into the sample jar. Rinse the cup with 95percent EtOH several times, adding the rinse to the sample jar. Hypsometric Pooling Rationale and Definition In March 1986 the LTER Zooplankton Group decided to pool the discrete depth Schindler Patalas trap samples into one pooled sample per lake-date for counting. Counting pooled samples rather than all of the depth samples reduces the time to produce zooplankton count data. The group hoped to count pooled samples from the entire backlog of uncounted samples and eventually to count samples shortly after collection. Samples are pooled considering lake hypsometry and, therefore, represent the entire lake. Previously, unpooled samples (2-9 samples per lake-date) or samples pooled considering only a water column were counted. Hypsometric pooling allows us to consider the zooplankton community as representing the entire lake, as our other limnological methods do, instead of just a column of water. Lake hypsometry is a three dimensional image of a lake or basin. In a simplified example of hypsometry, a lake is similar to a cone filled with water. If the cone were divided into three layers by two equidistant horizontal planes, the volumes in those layers would be very different from each other. The uppermost layer would contain the most water. Similarly, the upper depths typically contain most of the volume of a lake. Pooling is the creation of a new sample from subsamples of the Schindler trap samples collected from one lake-date. The volume of each subsample used to make the pooled sample reflects the depth range the sample represents and the volume of water that range represents relative to the entire lake volume. Samples pooled in this manner are called HP samples, for hypsometrically pooled, and are referred to as volume weighted because the volume of lake water each depth sample represents determines the subsample size. In sum, the advantages to this method of pooling are quicker turnover time and representation of the entire lake in a volume weighted fashion. Disadvantages of this method include the time required to pool subsamples, errors introduced during pooling, and the loss of more specific depth information. Pooling Procedure Allow the sample jars to air dry for a day or two. Weigh the Wisconsin Net sample and record the weight on the Volume by Weight form. Mark the liquid level on the jar with a Sharpee brand permanent marker. Add 95percent EtOH to each Schindler trap sample to bring liquid volume up to a weight of 105g, measured by weighing the sample jar with lid on the balance. If sample jars already contain more than 105g of liquid, allow some of the volume to evaporate in the hood, and then bring up to 105g. Record the final weight of jar plus sample plus EtOH on the Volume by Weight form. Calculate the subsample volumes, called target volumes, using the hypsometric table for each lake. Record these volumes on the Volume by Weight form. Mix the first sample gently and thoroughly by tilting the jar from side to side. Measure the target volume into a plastic graduated cylinder. Pour the subsample quickly and smoothly because the plankton settle out quite rapidly. Choose the smallest size graduated cylinder that can measure the target volume in one aliquot. Add the subsample to the labeled HP jar. Repeat with all other depth samples. When all of the subsamples have been added to the HP sample, rinse each graduated cylinder into the HP jar with several small volumes of EtOH. Place the HP sample in the hood to evaporate the excess volume of EtOH. The final weight of the HP sample should be 105g. Mark the liquid level on the jar with a Sharpee brand permanent marker. If the samples are from the August quarterly, pour the remainder of each Schindler sample into a labeled jar for archival. For all other sample dates, discard what is left of the Schindler samples. Rinse and air dry the field sample jars. Sample Storage and Record Keeping Store samples in cardboard records boxes obtained from UW Stores, storing samples from each lake in a separate box. Approximately one year of samples will fit in one box. Fill out the forms for each sample and sample box, as noted below. Box Inventory Form: A record of box contents. It remains in the sample storage box. Volume by Weight Form: A record of samples collected for any one lake-date and their volumes, storage box number, and history of sample usage. Filed in 3-ring binders, one copy at the Zoology Museum and one copy with Corinna Gries. Samples Stored Form: A record of all samples collected and storage box number for each. Current forms are kept in a binder at Trout Lake; archived forms are kept in the UW Zoology Museum. The data are eventually entered into the electronic LTER Museum Catalog. When boxes become full, check the contents against the Inventory Form and Samples Stored Form, and transfer them to the sample storage room in the garage. LTER samples and related paperwork are eventually transferred to the Zoology Museum at UW-Madison. Zooplankton Counting Before removing a subsample from any zooplankton sample jar, weigh the sample to check for evaporation. If the weight is within 0.1 gram of the last weight recorded on the Volume by Weight sheet, no fluid replacement is necessary. If the weight is more than 0.1 gram low, add 95percent EtOH to the sample to bring it up to the correct volume. Mix the sample well by turning the jar on its side and tipping back and forth gently. We use a Hensen-Stempel pipet with a 5-ml plunger for subsampling zooplankton samples. After mixing the sample, take the subsample as quickly as possible to avoid biasing the subsample as organisms begin to sink. There should be no air bubbles inside the Hensen-Stempel pipet. If there are, replace the subsample into the jar, completely dry the pipet, and begin again with the mixing. When you have a bubble-free subsample, dry the outside of the pipet and dispense the subsample into a cup with 53µ mesh bottom. Rinse the pipet into the cup with RO water, and continue rinsing the sample in the cup, washing the ethanol out of the sample through the mesh. Rinse the subsample into the counting tray with RO water, washing the mesh thoroughly to transfer all organisms into the tray. After removing subsample(s) from the jar, weigh the sample jar, and record this weight in a new column of the Volume by Weight form. Record the balance used, your initials, and the date at the top of the column, and add a column header such as Column C minus subsamples removed for counting . Do not put the subsample back into the sample jar after counting. Mark the new liquid level on the jar with a permanent marker. Replace the sample jar into the proper storage box. Count copepods and cladocerans first, identifying individuals to species wherever possible, and staging all copepodids. Measure a subset of each species. Then add a few drops of Lugol s solution to the subsample to stain it, and count the rotifers and nauplii. Count two subsamples for copepods and cladocerans. Count one subsample for rotifers and nauplii. If there are less than 100 of the dominant rotifer in one subsample, count a second subsample for rotifers and nauplii. Add milli-RO water to the tray as necessary to keep the surface of the subsample level. If the surface becomes concave as the subsample evaporates, it is difficult to focus clearly, and measurements may become distorted. Count all eggs attached to any species. For copepods and cladocerans, keep track of the number of individuals with eggs as well as the total number of eggs. Total number of eggs is sufficient for rotifers. Measurements are done as follows: Measure copepods from the tip of the head to the end of the urosome, excluding the caudal rami. Measure cladocerans from the tip of the head to the posterior of the carapace, excluding tailspine. However, measure helmeted Daphnia species from the anterior edge of the eye to the posterior of the carapace. Rotifers are not routinely measured, but where they have been, the total body length excluding spines is measured. Body width rather than length is measured for Conochilus, Conochiloides, and Collotheca. Describe, measure, and draw any unknown species on a separate sheet of paper. If possible, take a photograph of the unknown.</p>All records from 1981-1989 were modified in March 2015 to correct an error in how density had been calculated. Density values in many cases are significantly reduced. Densities are contained in three database tables. The original data is in dbmaker.zoop_raw; an intermediate table is dbmaker.zoop_all_density; and the final table (the one this website extracts density from) is dbmaker.zoop_allnl_summary_snap. Density values are modified from the original to final tables as they are summed or averaged over other variables (sample depth, replicate, and sex stage). The table that was corrected in this case is dbmaker.zoop_all_density. The correction algorithm is as follows: Records from dbmaker.zoop_raw are first grouped to isolate each unique 3-tuple of lake, sample date, and species. Each group is subsequently treated independently. Multiplying the fields 'hp_factor' and 'no_per_l' results in a density value for that record. Density values are then summed within each unique replicate (sex stage is what varies within a replicate). These resultant sums are then averaged over all replicates, giving a density value for each lake, sample date, species, and depth combination. The result is written into field 'number_per_liter' in table dbmaker.zoop_all_density. Densities are subsequently summed over depth before being provided via the website. Records after 1989 were already valid and did not require any modification.</p>Taxonomic resolution: A genus-only designation could mean a different species than the otherwise named species, or it could mean that the person counting only identified it to genus.&nbsp; Within one sample (same lake and date) it may be assumed that a genus only individual is a different species than other SameGenus/Namedspecies in the same count.</p>
Short Name
NTLPL03
Version Number
37
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