US Long-Term Ecological Research Network

Lake Mendota, Wisconsin, USA, Zebra Mussel Body Size and Biomass Biometrics 2018

Abstract
We sampled 98 individuals of the zebra mussel (Dreissena polymorpha) population of Lake Mendota from many littoral zone sites in 2018 to create biometric relationships between several metrics of body size and several metrics of biomass, including length, width, height, living weight, wet weight, dry weight, shell weight, shell-free dry weight, and ash-free dry weight. We selected individuals to span a wide range of body sizes and found strong relationships between most combinations of body size and biomass metrics.<br/>
Dataset ID
395
Date Range
-
LTER Keywords
Methods
In the laboratory, three measurements of body size and seven measurements of biomass were captured. First, any foreign material found adhering to the external surface of specimens was completely removed. Body size directional measurements of shell length (L), width (W), and height (H) were recorded for every specimen with the aid of callipers (0.01 mm). Following this, any excess water was removed from surfaces by drying the external shell with tissue paper. Further, using a scalpel blade and tweezers, excess water was removed from the mantle cavity by gently forcing bivalves to gape, taking care not to cut the adductor muscle or damage tissues. Using high-resolution scales, living-weight (LW) was obtained for each specimen. Then each specimen was fully opened, which in most cases involved cutting of the adductor muscles. To remove additional fluid from the mantle and other cavities, each specimen was then placed with the valve gape (flesh) facing downwards onto absorbent tissue, for ~5-10 minutes. A wet-weight (WW) was obtained for each specimen. Following this, the soft tissue was dissected from the shell, then both soft tissue and shell were dried together within an oven (60-72 degreeC) for ~48 hrs, or until they reached a constant weight. Specimens were cooled to room temperature in a desiccator before final weighing. A combined dry-weight (DW) was recorded, as were weights for the soft tissue and shell separately, i.e. shell free dry-weight (SFDW) and dry shell-weight (SW), respectively. Following the establishment of SW, SFDW was calculated subtracting SW from the total DW (i.e. SFDW = DW–SW). To obtain an ash-weight (AW), the soft and hard tissue structures of specimens were incinerated (500–550 degreeC) together within a muffle furnace for 4–6 hrs. In all cases, the ash free dry-weight (AFDW) was then calculated for the entire specimen (soft tissue and shell) by subtracting the AW from DW, i.e. AFDW = DW–AW.<br/>
Version Number
1

Lake Mendota, Wisconsin, USA, Zebra Mussel Veliger Water Column Density 2016-2019

Abstract
We sampled veliger (larval stage) zebra mussels (Dreissena polymorpha) from 2016-2019.
Zebra mussels are invasive in Lake Mendota and were first detected in November 2015. Samples
were taken at three different sites on Lake Mendota from June to August in 2016, and from
June to November in 2018-2019, using a 0.5 m diameter, 64 micrometer mesh size plankton net
for an 8 m depth tow. This dataset complements adult zebra mussel, zoobenthos, and
phytobenthos data collected during the same time period, for which data is also archived
with EDI.
Core Areas
Dataset ID
392
Data Sources
Date Range
-
Methods
We sampled larval zebra mussels (veligers) using a 64 microm mesh, 0.5 m diameter
plankton net and stored them in 80% ethanol in 200 mL containers at 25degree C for 0-12
weeks until processing. At each site we performed triplicate 8 m depth plankton tows by
pulling a net from 2 m above the lake bottom at the 10 m depth sites of transects A-C
developed for adult zebra mussel collection. We collected samples approximately every 14
days from June to August in 2016, and June to November in 2017-2019. During fall
sampling, poor weather conditions occasionally limited the number of sites or replicates
collected. We also sampled veligers biweekly in 2019 but reduced sampling to one
replicate per site and only sampled at one site after September. Because veligers are
small and difficult to see, enumeration was time consuming. <br/>We sampled larval zebra mussels (veligers) using a 64 microm mesh, 0.5 m diameter
plankton net and stored them in 80% ethanol in 200 mL containers at 25degree C for 0-12
weeks until processing. At each site we performed triplicate 8 m depth plankton tows by
pulling a net from 2 m above the lake bottom at the 10 m depth sites of transects A-C
developed for adult zebra mussel collection. We collected samples approximately every 14
days from June to August in 2016, and June to November in 2017-2019. During fall
sampling, poor weather conditions occasionally limited the number of sites or replicates
collected. We also sampled veligers biweekly in 2019 but reduced sampling to one
replicate per site and only sampled at one site after September. Because veligers are
small and difficult to see, enumeration was time consuming. <br/>We sampled larval zebra mussels (veligers) using a 64 microm mesh, 0.5 m diameter
plankton net and stored them in 80% ethanol in 200 mL containers at 25degree C for 0-12
weeks until processing. At each site we performed triplicate 8 m depth plankton tows by
pulling a net from 2 m above the lake bottom at the 10 m depth sites of transects A-C
developed for adult zebra mussel collection. We collected samples approximately every 14
days from June to August in 2016, and June to November in 2017-2019. During fall
sampling, poor weather conditions occasionally limited the number of sites or replicates
collected. We also sampled veligers biweekly in 2019 but reduced sampling to one
replicate per site and only sampled at one site after September. Because veligers are
small and difficult to see, enumeration was time consuming.
Version Number
1

North Temperate Lakes LTER: Trout Lake Spiny Water Flea 2014 - present

Abstract
Beginning in 2014, 30 meter vertical tows with a special zooplankton net were collected in Trout Lake specifically for the invasive Bythotrephes longimanus (spiny water flea). The net has a 400 micrometer mesh with a 0.5 meter diameter opening. Individuals are simply counted, and density is determined to be the number of individuals divided by the total water volume of each tow.
Additional Information
Related data set: North Temperate Lakes LTER: Zooplankton - Trout Lake Area 1992 - current (37)
Core Areas
Dataset ID
389
Date Range
-
Publication Date
Version Number
1

North Temperate Lakes LTER Regional Survey Zooplankton 2015 - current

Abstract
The Northern Highlands Lake District (NHLD) is one of the few regions in the world with periodic comprehensive water chemistry data from hundreds of lakes spanning almost a century. Birge and Juday directed the first comprehensive assessment of water chemistry in the NHLD, sampling more than 600 lakes in the 1920s and 30s. These surveys have been repeated by various agencies and we now have data from the 1920s (UW), 1960s (WDNR), 1970s (EPA), 1980s (EPA), 1990s (EPA), and 2000s (NTL). The 28 lakes sampled as part of the Regional Lake Survey have been sampled by at least four of these regional surveys including the 1920s Birge and Juday sampling efforts. These 28 lakes were selected to represent a gradient of landscape position and shoreline development, both of which are important factors influencing social and ecological dynamics of lakes in the NHLD. This long-term regional dataset will lead to a greater understanding of whether and how large-scale drivers such as climate change and variability, lakeshore residential development, introductions of invasive species, or forest management have altered regional water chemistry. Zooplankton samples were taken at approximately the deepest part of each lake, via a vertical tow with a Wisconsin net. Count of individuals and presence absence data for all lakes in study region are provided here.
Contact
Core Areas
Dataset ID
381
Date Range
-
Maintenance
ongoing
Methods
Each zooplankton sample was taken at approximately the deepest part of each lake, via a vertical tow with a Wisconsin net (20cm diameter mouth, 80µ mesh) lowered to 1 meter above the bottom of a lake and then pulled up slowly at a rate of about 3 seconds per meter. Contents of the net were preserved in 4-oz jars with 95% ethanol. One sample was taken from each lake. Samples were collected by the Regional Lakes summer sampling crew in June 2015.
Version Number
1

North Temperate Lakes LTER Zooplankton conversion formulas length to biomass

Abstract
Formulas for calculating zooplankton biomass based on measured length for species encountered in NTL's northern lakes. Formulas are either based on literature reports or measurements in particular research lakes.
Core Areas
Dataset ID
376
LTER Keywords
Maintenance
completed
Methods
formulas are based on data in literature or were determined in samples from research lakes:

Culver D.A. et.al. 1985. Can. J. Fish. Aquat. Sci. Vol 42, 1380-1390.
Biomass of freshwater crustacean zooplankton from length-weight regressions.

Downing, John A. and Frank H. rigler. 1984.
A manual on methods for the assessment of secondary productivity in fresh waters. Second edition.

Dumont, H.J., I. Van de Velde and S. Dumont. Ref??
The dry weight estimate of biomass in a selection of cladocera, copepoda and rotifera from the plankton, periphyton and benthos of continental waters.

Hawkins, Bethany E. and M.S. Evans. 1979. J.Great Lakes Res. 5(3-4):256-263
Seasonal cycles of zooplankton biomass in southeastern Lake Michigan

Lawrence, S.G., D.F. Malley, W.J. Findlay, M.A. MacIver and I.L. Delbaere. 1987. Can J. Fish. Aquat. Sci. 44: 264-274.
Methods for estimating dry weight of freshwater planktonic crustaceans from measures of length and shape.

Pace M.L. and J.D. Orcutt. 1981. Limnol. Oceanogr. 26(5), 822-830.
The relative importance of protozoans, rotifers, and crustaceans in a freshwater zooplankton community.

Yan N.D. and G.L. Mackie. 1987. Can. J. Fish. Aquat. Sci. Vol 44, 382-389.
Improved estimation of the dry weight of Holopedium gibberum using clutch size, a body fat index, and lake water total phosphorus concentration.

Ruttner-Kolisko A. 1977. Arch. Hydrobiol. Beih. Ergebn. Limnol. 8, 71-76.
Suggestions for biomass calculations of plankton rotifers.
Version Number
1

Little Rock Lake Experiment at North Temperate Lakes LTER: Zooplankton length 1988 - 1998

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 average length of organisms for each species.
Core Areas
Dataset ID
375
Date Range
-
Maintenance
completed
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.
Version Number
1

Cascade project at North Temperate Lakes LTER - Daily data for key variables in whole lake experiments on early warnings of critical transitions, Paul and Peter Lakes, 2008-2011

Abstract
Peter Lake's food web was altered by adding largemouth bass at a slow rate while monitoring key food web constituents including littoral minnow abundance indexed as catch per trap per hour, zooplankton biomass, and concentration of chlorophyll a. Paul Lake was manipulated and the same variables were measured there.
In Peter Lake, we expected littoral catch of minnows to first increase as minnows moved into the littoral zone due to the threat of bass predation and then decrease due to bass predation. We expected zooplankton biomass to increase as minnows moved into the littoral zone. We expected chlorophyll to decrease due to increased grazing by zooplankton. We expected that variance and autocorrelation of chlorophyll would increase as the food web passed a critical transition.
We expected that the time series in Paul Lake would represent the normal variability of an unmanipulated lake
Dataset ID
374
Date Range
-
Methods
Primary publications that provide more information about taxa, methods, and data are:
Carpenter, S.R., J.J. Cole, M.L. Pace, R.D. Batt, W.A. Brock, T. Cline, J. Coloso, J.R. Hodgson, J.F. Kitchell, D.A. Seekell, L. Smith and B. Weidel. 2011. Early warnings of regime shifts: A whole-ecosystem experiment. Science 332: 1079-1082.
Cline, T.J., D. A. Seekell, S. R. Carpenter, M. L. Pace, J. R. Hodgson, J. F. Kitchell, and B. C. Weidel 2014. Early warnings of regime shifts: evaluation of spatial indicators from a whole-ecosystem experiment. Ecosphere 5:art102. http://dx.doi.org/10.1890/ES13-00398.1
Pace, M.L., S.R. Carpenter, R.A. Johnson and J. T. Kurzweil. 2013. Zooplankton provide early warnings of a regime shift in a whole-lake manipulation. Limnology and Oceanography 58: 525-532.
For an explanation of our rationale and expected results see:
Carpenter, S. R., Brock, W. A., Cole, J. J., Kitchell, J. F., & Pace, M. L. 2008. Leading indicators of trophic cascades. Ecology Letters, 11(2), 128-138. doi:DOI 10.1111/j.1461-0248.2007.01131.x
Version Number
2

Cascade Project at North Temperate Lakes LTER Core Data Zooplankton 1984 - 2016

Abstract
Zooplankton data from 1984-2016. Sampled approximately weekly with two net hauls through the water column (30 cm diameter net, 80 um mesh). There have been over eight zooplankton counters during this period, so species-level identifications (TAX, below) are not as consistent as those for some of the other datasets. Sampling Frequency: varies; Number of sites: 8
Core Areas
Dataset ID
355
Date Range
-
Maintenance
completed
Methods
Sampling:
Zooplankton were sampled approximately weekly with two net hauls through the water column (30 cm diameter net, 80 um mesh). Tows were taken at standard depths for almost all years. The standard depths are as follows: Peter, East Long, West Long, Crampton and Tuesday Lakes: 12m, Paul Lake: 8m, Ward Lake: 6m; exceptions are: for 2012 and beyond Tuesday Lake was sampled at 10m, Peter was sampled at 10m from 1984-1986, Paul was sampled at 7.5m in 1995. Samples were preserved with cold sugared formalin or Lugol's solution.
Version Number
16

North Temperate Lakes LTER Bythotrephes longimanus spiny water flea population monitoring in Wisconsin and Minnesota 2009 - 2014

Abstract
Three data tables are included describing population dynamics for Bythotrephes longimanus, spiny water flea, in Southern Wisconsin during invasion. General monitor took place in Lake Mendota, Lake Monona, Lake Waubesa, Lake Kegonsa, Stormy Lake, Gile Flowage, Lake Gogebic.Accompanying Bythotrephes morphological measurements from Lake Mendota monitoring efforts in 2011 and 2012. Included are individual measurements of body morphology and reproductive status for ~2,500 <em>Bythotrephes </em>collected from Lake Mendota in 2011 and 2012.Sediment cores from Lake Mendota were analyzed for spiny water flea evidence with age of sediment estimated.
Contact
Core Areas
Dataset ID
342
Date Range
-
Maintenance
complete
Methods
general monitoring for spiny water flea:
The dataset contains collected Bythotrephes longimanus monitoring efforts from 8 invaded lakes in Wisconsin that took place over the course of 2009 through 2014 using a zooplankton net. Monitoring efforts were conducted to 1) obtain more accurate estimates of Bythotrephes densities using a more appropriately sized net (50-cm diameter over 30-cm diameter) and 2) obtain detailed demographic measurements of Bythotrephes morphology and reproduction in each lake. Here only Bythotrephes densities are included.
The majority of samples occurred at a lakes deep hole with a 50-cm diameter and 150-micron mesh zooplankton net. Nets are lowered to 2 m off of the lake bottom before being towed to the surface. Samples are processed in their entirety
Exceptions to this are those at sites containing “LTER” (e.g., site IDs LTER-DH and LTER-MB) in their ID which were samples taken according to the Southern Lakes LTER zooplankton collection protocol with a 30-cm and 83-micron mesh. Other exceptions include sites outside the deep hole of the lake (site ID 5m = 5m lake depth north of the Center for Limnology on Lake Mendota; CFL = 15m lake depth north of the Center for Limnology; DH = deep hole but specific to Lake Mendota; MB = 15m lake depth southwest of Maple Bluffs in Madison on Lake Mendota; MO.5m = a 5m lake depth site in Lake Monona; MO.Y = 5m lake depth site at the mouth of the Yahara River on Lake Monona; TL = 15m lake depth west of Tenney Locks in Madison on Lake Mendota; WS = 15m site in northwestern basin of Lake Mendota, east of Picnic Point; WP = 5m site south of Warner Park on Lake Mendota). Several tows were taken using a 200m oblique (i.e., horizontal) net tow with the 50-cm diameter net (DH-ObliqueTow). Efforts in Southern Wisconsin were led by Jake Walsh while efforts in Northern Wisconsin were led by Carol Warden (site ID = CW), Pam Montz (site ID = PM), Sam Christel (site ID = SC), Sam Oliver (site ID = SM), as well as a researcher with initials (site ID) “EM”.
Version Number
8

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.
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