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.

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.

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.

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.

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.

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.

Methods

BOOM SHOCKING1987:A standard WDNR electrofishing boat was used on Lake Mendota set at 300 volts and 2.5 amps (mean) DC, with a 20 % duty cycle and 60 pulses per second. On all sampling dates two people netted fish, the total electrofishing crew was three people. Shocking was divided into stations. For each station, the actual starting and ending time was recorded. Starting and ending points of each station were plotted on a nap. A 7.5 minute topographic map (published 1983) and a cartometer was used to develop a standardized shoreline mileage numbering scheme. Starting at the Yahara River outlet at Tenney Park and measuring counterclockwise, the shoreline was numbered according to the number of miles from the outlet. The length of shoreline shocked for each station was determined using the same maps. The objectives of the fall 1987 electrofishing was: to gather CPE data for comparison with previous surveys of the lake; develop a database for relating fall electroshocker CPE to predator density; collect fall predator diet data; make mark-recapture population estimates of YOY predators; and determine year-class-strength of some nonpredators (yellow perch, yellow bass, and white bass).1993: Electrofishing was used to continue marking largemouth and smallmouth bass (because of low CPE in fyke nets), to recapture fish marked in fyke netting, and to mark and recapture walleyes ( less than 11.0 in.) on Lake Mendota. Four person crews electrofished after sunset from May 05 to June 03, 1993. A standard WDNR electrofishing boat was used, set at about 300 volts and 15.0 amps (mean) DC, with a 20 % duty cycle at 60 pulses per second. On all sampling dates two people netted fish; thus, CPE data are given as catch per two netter hour or mile. Shocking was divided into stations. For each station the actual starting and ending time and the generator s meter times was recorded. Starting and ending points of each station were plotted on a map. 7.5 minute topographic maps (published in 1983) were used in addition to a cartometer to develop a standardized shoreline mileage numbering scheme. Starting at the Yahara River outlet at Tenney Park and measuring counterclockwise the shoreline was numbered according to the number of miles from the outlet. The length of shoreline shocked for each station was determined using these maps. The 4 person electroshocker crews were used again from September 20 to October 19. Fall shocking had several objectives: to gather CPE data for comparison with previous surveys of the lake; develop a database for relating fall electroshocker CPE to piscivore density; and make mark recapture population estimates of young of year (YOY) piscivores.1997:5/13/1997-5/20/1997: Electrofishing was completed at night on lakes: Mendota, Monona, and Waubesa. A standard WDNR electrofishing boat was used, set from 320-420 volts and 16-22 amps DC, with a 20 % duty cycle at 50 pulses per second. Two netters were used for each shocking event. At a particular station, starting and ending times where shocking took place were recorded. The location of the designated shocking stations is unknown.9/23/1997-10/14/1997: Electrofishing was completed at night on Mendota, Monona, Waubesa, and Wingra. A standard WDNR electrofishing boat was used, set from 315-400 volts and 16-24 amps DC, with a 20% duty cycle at 60 pulses per second. Two netters were used for each shocking event. Starting and ending time at each shocking station was listed. The location of the designated shocking stations is unknown.1998:Electrofishing was completed at night on Mendota, Monona, Wingra, and Waubesa from 5/12/1998- 10/28/1998. A standard WDNR electrofishing boat was used, set from 240-410 volts and 15-22 amps DC, with a 20% duty cycle at 50-100 pulses per second. Two netters were used for each shocking event. Starting and ending time at each shocking station was listed. The location of the designated shocking stations is unknown. FYKE NETTING1987:Fyke nets were fished daily from March 17 to April 24, 1987 on Lake Mendota. The nets were constructed of 1.25 inch (stretch) mesh with a lead length of 50 ft. (a few 25 ft. leads were used). The hoop diameter was 3 ft. and the frame measured 3 ft. by 6 ft. Total length of the net was 28 ft. plus the lead length. Nets were set in 48 unknown locations. Initially, effort was concentrated around traditional northern pike spawning sites (Cherokee Marsh, Sixmile Creek, Pheasant Branch Creek, and University Bay). As northern pike catch-per-effort (CPE) declined some nets were moved onto rocky shorelines of the lake to capture walleyes. All adult predators (northern pike, hybrid muskie, largemouth and smallmouth bass, walleye, gar, bowfin, and channel catfish) captured were tagged and scale sampled. Measurements on non-predator species captured in fyke nets were made one day per week. This sampling was used to index size structure and abundance, and to collect age and growth data. In each net, total length and weight of 20 fish of each species caught was measured, and the remaining caught were counted.1993:Same methods as 1987, except fyke nets were fished from 4/8/1993-4/29/1993 on Lake Mendota. The 1993 fyke net data also specifies the &ldquo;mile&rdquo; at which the fyke net was set. This is defined as the number of miles from the outlet of the Yahara River at Tenney Park, moving counterclockwise around the lake. In addition, abundance and lengths of non-gamefish species captured in fyke nets were recorded one day per week. Six nets were randomly selected to sample for non-gamefish data. This sampling was used to index size structure and abundance, and to collect age and growth data. In each randomly selected net, total length and weight was measured for 20 fish of each species, and the remaining caught were counted.1998:There is no formal documentation for the exact methods used for fyke netting from 3/3/1998-8/12/1998 on Lake Mendota. However, given that the data is similar to data collected in 1987 and 1993 it is speculated that the same methods were used.MINI-FYKE NETTING1989:There is no formal documentation for the exact methods used for mini-fyke netting on Lake Mendota and Lake Monona from 7/26/1989-8/25/1989. However, given that the data is similar to data collected from 1990-1993 it is speculated that the same methods were used. In the sampling year of 1989, mini-fyke nets were placed at 22 different unknown stations.1990-1993: Mini-fyke nets were fished on Lake Mendota and Lake Monona during July-September at 20, 29, 13, and 15 sites per month during 1990, 1991, 1992, and 1993, respectively to estimate year-class strength, relative abundance, and size structure of fishes in the littoral zone. Nets were constructed with 3/16 in. mesh, 2 ft. diameter hoops, 2 ft. x 3 ft. frame, and a 25 ft. lead. Sites were comparable to seine sites used in previous surveys. Sites included a variety of substrate types and macrophyte densities. To exclude turtles and large piscivores from minifyke nets, some nets were constructed with approximately 2 in. by 2 in. mesh at the entrance to the net. Thus, mini-fyke net data are most accurate for YOY fishes, and should not be used to make inferences about fishes larger than the exclusion mesh size. 1997:There is no formal documentation for the mini-fyke methods which were used on Lake Waubesa and Lake Wingra from 9/16/1997-9/18/1997. However, given that the data is similar to data collected in 1989, and 1990-1993, it is speculated that the methods used during 1997 are the same. SEINE NETTING1989, 1993: Monthly shoreline seining surveys were conducted on Lake Mendota and Lake Monona during June through September to estimate year class-strength, relative abundance, and size structure of the littoral zone fish community. Twenty sites were identified based on previous studies. Sites included a variety of substrate types and macrophyte densities. Seine hauls were made with a 25ft bag seine with 1/8 inch mesh pulled perpendicular to shore starting from a depth of 1 m. Twenty fish of each species were measured from each haul and any additional fish were counted. Gill Netting (1993)Experimental gill nets were fished in weekly periods during June through August, 1993. Gill nets were used to capture piscivores for population estimates of fish marked in fyke nets. All nets were constructed of five 2.5-4.5 in. mesh panels, and were 125 ft. long. Nets set in water shallower than 10 ft. were 3ft. high or less; all others were 6ft. high or less. Sampling locations were selected randomly from up to three strata: 1) offshore reef sets, 2) inshore sets, 6.0-9.9 ft. deep, and 3) mid-depth sets, 10-29.9 ft. deep. The exact location at which the gill nets were set on the lake is unknown because the latitude and longitude values which were recorded by the WDNR are invalid. Temperature and dissolved oxygen profiles were used to monitor the development of the thermocline and guide net placement during July and August. After the thermocline was established nets were set out to the 30 ft. contour or to the maximum depth with dissolved oxygen greater than 2 ppm. Walleye Age: Scale and Spine Analysis (1987) Scales were taken from walleye that were shocked during the fall of 1987 electrofishing events on Lake Mendota. Scales were taken from 10 fish per one-inch length increment. The scales were removed from behind the left pectoral fin, and from the nape on the left side on esocids. In addition, the second dorsal spine was removed from 10 walleyes per sex and inch increment (to age and compare with scale ages for fish over 20 inches). CREEL SURVEYS1989:Fishing pressure, catch rates, harvest, and exploitation rates were estimated from a randomized, access-point creel survey. The schedule was stratified into weekday and weekend/holiday day types. Shifts were selected randomly and were either 07:00-15:00 h or 15:00-23:00 h. In addition, two 23:00-03:00 h shifts and two 03:00-07:00 h shifts were sampled per month to estimate the same parameters during night time hours. During the ice fishing season (January-February) 22 access points around Lake Mendota and upstream to the Highway 113 bridge were sampled. The clerk counted the number of anglers starting and completing trips during the scheduled stop at each access point. During openwater (March-December) 13 access points were sampled; 10 were boat ramps and 3 were popular shore fishing sites<strong>. </strong>At each of these sites, an instantaneous count of shore anglers was made upon arrival at the site, continuous counts of anglers starting and completing trips at public and private access points were made. Boat occupants and ice fishing anglers were only interviewed if they were completing a trip. Both complete and incomplete interviews were made of shore anglers. Number caught and number kept of each species, and percent of time seeking a particular species were recorded. All predators possessed by anglers were measured, weighed, and inspected for finclips and tags. We measured a random sample of at least 20 fish of each non-predator species per day.1990-1993: Same as 1989, except 23 access points were used during the ice fishing season. In addition, 13 access points were sampled during the openwater (May-December) season; 9 sites were boat ramps and 4 sites were popular shore fishing sites. 1994-1999: No formal documentation exists, but given the similarity in the data and consistency through the years; it is speculated tha tthe methods are the same.