Methods

Mussels that were collected for glycogen analysis were immediately placed on ice to preserve the tissue for later analysis (following Dunn and Ellis, 2005). In the lab, individuals were sexed via external shell morphology, dried in a drying oven to a constant mass, weighed, then approximately 10mg of foot tissue was collected and run for glycogen analysis (following Naimo et al. 1998). For mussels that were sampled for metabolomics analysis, in the field, immediately after collection, approximately 200uL of hemolymph was drawn from the anterior adductor muscle using a 27–29-gauge syringe, stored in a 2mL cryotube, and flash frozen in liquid nitrogen. Unionids were then scraped of all zebra mussels if present and returned to the benthos. All zebra mussels greater than length 5mm were counted and measured along the major axis of the shell. Cumulative dry weight of all zebra mussels was estimated using a standard conversion of length to dry weight for zebra mussels (Coughlan et al. 2021). Zebra mussel loads were calculated as total estimated zebra mussel dry mass divided by the total dry mass of the unionid soft tissue. Hemolymph samples were stored at -80˚C until they were shipped on dry ice to West Coast Metabolomics Center at UC-Davis for non-targeted analysis of primary metabolites. Metabolites were identified using gas chromatography time-of-flight mass spectroscopy (GC-TOF MS and their peak heights were normalized to the sum peak height of annotated compounds (mTIC-normalization) by the West Coast Metabolomics Center. Each metabolite was independently normalized to zero and standardized to one standard deviation.<br/>

Methods

Phytoplankton samples were obtained from the epilimnion and hypolimnion by
slowly lowering weighted Tygon tubing through the water column to a depth of 7 m, such
that the tubing was filled with a representative water column sample. Based on the inner
diameter of the tubing, 205 mL of water was pumped from the tubing for the epilimnion
sample. Next, 267 mL of water was pumped from the tubing for the hypolimnion sample. Each
sample was collected into a 250 mL amber bottle that contained 2 mL of Lugol’s solution.
Phytoplankton samples were pooled by sampling date and sent to Phycotech Inc. (St. Joseph
MI, USA) for phytoplankton identification, and concentration and biovolume
quantification.<br/>

Methods

Our study lake, South Sparkling Bog (SSB) (46.003°N, 89.705°W), is a bog lake
located in Vilas County in Northern Wisconsin. South Sparkling Bog is a dystrophic,
dimictic lake with a maximum depth of 8 m, a mean depth of 3.6 m, and a surface area of
0.44 ha. South Sparkling Bog is surrounded by a sphagnum bog mat and has no shoreline
development. During the winters of 2019-2020 and 2020-2021, snow was removed from the
surface of South Sparkling Bog following any snow accumulation event. Removal was
conducted via a snowplow attached to the front of an ARGO all-terrain vehicle and a
snowblower. The winter of 2018-2019 served as a reference year, and snow was not removed
from South Sparkling Bog’s surface. While ice cover persisted, plankton samples were
collected at the deep spot for each lake during on a biweekly-to-monthly basis each
winter. On each sampling date, one integrated zooplankton tow was taken at a depth of 0-7
m using a 56 µm mesh Wisconsin net. All zooplankton samples were collected into glass
sample jars, preserved in 90% ethanol, and saved for laboratory analysis. In the lab,
zooplankton samples were filtered through 53 µm mesh and diluted to a known volume, and
three sub-sample replicates were taken using a 1 mL Hensen-Stempel pipette. Sub-sample
replicates were counted to at least 100 individuals, otherwise the entire sample was
quantified. Sub-sample data was then converted to the known diluted volume and finally
converted to total filtered volume (from the integrated tow sample) to estimate density
(individuals L-1). Zooplankton samples were processed using a Leica M8Z dissecting scope
and Leica imaging software. Replicate subsamples were averaged to estimate total abundance
and density, and average lengths (mm) for each sample taxa were calculated from measures
of the first 30 taxa found within a sample date.<br/>

Methods

Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>Nutrients were added to Peter (2013-2015, 2019) and Tuesday (2013-2015)
lakes to cause algal blooms. Details on nutrient additions (start/end dates,
loading rates, N:P ratios) are described in Buelo et al. 2022 (Ecological
Applications, link below), Wilkinson et al. 2018 (Ecological Monographs 88:
188-203), and Pace et al. 2017 (Proceedings of the National Academy of
Sciences USA 114: 352-357). These publications including supplements should
be consulted for details. These lakes have been used for whole-ecosystem
experiments over the past decades; see Carpenter and Pace 2018 (Limnology
and Oceanography Letters 3(6): 419-427) for an overview.<br/>

Methods

Study area
The experimental (McDermott Lake; 46.00299280, -90.16081610) and reference (Sandy Beach Lake (46.10614350, -89.97131020) systems are located in Iron County in Northern, Wisconsin. McDermott Lake has a surface area of 33.1 ha and a mean depth of 3.0 m, while Sandy Beach Lake has a surface area of 44.5 ha and a mean depth of 2.1 m. Maximum depth in McDermott Lake is 5.7 m and in Sandy Beach Lake is 4.0 m. Both lakes include a variety of substrates (e.g., rock, gravel, and sand) and areas of submerged and emergent vegetation. At the start of the study, McDermott and Sandy Beach Lake fish communities were similar with high Centrarchidae spp. (i.e., centrarchid) abundances, few adult walleye, and a history of natural walleye recruitment. Other species present include yellow perch, northern pike, muskellunge, black bullhead, and white sucker (Table 1).
Fish sampling
Standardized surveys
From 2017-2020, we conducted fish population sampling on the experimental (McDermott) and reference (Sandy Beach) lakes. Every year of the study, we have conducted standardized monitoring surveys employing numerous sampling techniques to detect changes in the fish community relative to 2017-baseline information (Fig. 1). Sampling began immediately following ice-out (~mid-April) with the deployment of five fyke nets for one week. The fyky surveys served two purposes: 1) to serve as the capture method for marking walleye as part of the mark-recapture survey to attain a population estimate, and 2) to estimate other focal species (i.e., black crappie, yellow perch, muskellunge, northern pike) relative abundances. During these surveys, all collected walleye were measured (total length (TL); mm), sexed, checked for a Passive Integrated Responder (PIT) tag, and if one was not present, marked with a unique PIT tag. We also removed a dorsal spine sample for aging. Adult (mature) walleyes were defined as all fish 381 mm and all fish for which sex could be determined (regardless of length). Walleye of unknown sex &lt;381 mm were classified as juvenile (immature). McDermott and Sandy Beach Lakes have both had walleye population estimates previously conducted by the Wisconsin Department of Natural Resources (WDNR) therefore the goal was to mark 10% of the anticipated spawning population (based off of previous population estimates). Marking continued until the target number was reached or spent females began appearing in the fyke nets. Walleye were recaptured using an AC boat electrofishing survey within one week (typically 1-4 days) after netting and marking were completed. In each lake, the entire shoreline was sampled. All captured walleyes were measured and examined for marks. Based on electrofishing mark-recapture data, population estimates were calculated using the Chapman (1951) modification of the Petersen Estimator as:
N=((M+1)(C+1))/((R+1))
where N was the population estimate, M was the number of fish marked and released, C was the total number of fish captured and examined for marks in the recapture sample, and R was the total number of marked fish observed in C. The Chapman Modification method was used because it provides more accurate population estimates in cases when R is relatively small (Ricker 1975).

From early-May to mid-June, we sampled larval fishes using a 1,000m mesh conical ichthyoplankton net towed for five minutes immediately below the lake surface. Weekly samples were taken at night at five sampling locations in each lake. Each lake was divided into five quadrats and sites were established at a randomly selected nearshore (&lt;100 m of shore) location in each quadrat on each sampling date. Once selected, locations remained fixed throughout the study. Volume of water filtered during each tow was estimated using a General Oceanics© model 2030R flowmeter mounted in the center of the net frame. Samples were transferred to containers and stored in 90% ethanol. Collected fishes were identified to species according to Auer (1982) and enumerated.
In addition to the adult walleye population, we were interested in estimating the size of the adult largemouth bass population. We performed early summer (late-May) mark-recapture surveys using AC boat electrofishing (Wisconsin‐style; AC; 2.0–3.0 amps, 200–350 V, 25% duty cycle with two netters) to sample largemouth bass. Collected largemouth bass were measured, checked for a top caudal fin clip, and if not present, marked with a top caudal fin clip and released. Adult (mature) largemouth bass were defined as all fish 203 mm. We aimed to recapture 10% of the marked population. Largemouth bass are in very low abundance in Sandy Beach Lake therefore a population estimate was not possible. In McDermott Lake, due to the small population size of largemouth bass we completed multiple marking surveys from late-May to early June to achieve this recapture rate. From electrofishing mark-recapture information, population estimates were calculated using the Schnabel (1938) modification of the Lincoln-Petersen method:
where N was the population estimate, M was the number of fish marked and released in sample t, C was the number of fish captured in sample t, and R was the number of fish already marked when caught in sample t.
To obtain centrarchid population demographic data, current standardized WDNR surveys of inland lakes consist of early summer (water temperature range = 13.0–21.0°C) AC boat electrofishing surveys or mid‐summer (18.3–26.7°C) mini‐fyke net (Simonson et al. 2008). To encompass this range of water temperatures, we performed a combination of surveys starting at the end of May with an AC boat electrofishing survey. Then, fish were sampled once monthly when lake surface water temperatures were ≥13.0°C in both lakes (June–September). Both lakes were sampled during 1‐week each month using three gears (AC boat electrofishing, mini-fyke nets, cloverleaf traps). Lakes were sampled on consecutive nights in each 1‐week period but only one gear type was employed per night.
All gears sampled shallow shorelines (0–5 m from bank, depth ≤2 m) and were deployed in fixed locations following standard approaches (Bonar et al. 2009). Sampling locations were evenly distributed along the shoreline of the lake, and all gears were deployed in similar habitat types. Five 10‐min nighttime boat electrofishing (Wisconsin‐style; AC; 2.0–3.0 amps, 200–350 V, 25% duty cycle) transects were conducted using two dipnetters. Five mini‐fyke nets (0.9‐m × 0.61‐m frames, 3.2‐mm mesh [bar measure], 7.6‐m‐long lead, and a double throat) were deployed in areas where the net frames would be in 1.0–1.5 m of water, and leads were fixed onshore. Five cloverleaf traps (three lobed, height = 41 cm, 50 cm diameter, 6.0‐mm bar wire mesh with 12.7‐mm‐wide openings between lobes, and an attractant [liver]) were deployed in littoral habitats. Both mini‐fyke nets and cloverleaf traps were set in early afternoon, fished overnight, and retrieved the following afternoon (~24‐h soak time). All catches were standardized according to gear-specific effort. For boat electrofishing, catch per unit effort (CPUE) is presented as fish/hr. For mini-fyke nets and cloverleaf traps, CPUE is calculated as fish/net night or fish/trap night.
To quantify walleye recruitment in each lake, we employed multiple gears throughout the sampling season including micromesh gillnets, beach seines, and boat electrofishing. In late July/early August, we deployed four 46-m x 1.2-m gillnets with 0.95-cm bar mesh. Sampling locations were evenly distributed along the shoreline and locations were fixed each year. Gillnets were set at night and at depths ranging from 0-5 m. Set duration ranged 1-2 hours to minimize bycatch, thus catches were standardized to age-0 walleyes collected per 10 hours of soak time. In late August, we pulled .24-m long beach seines with 0.64-cm mesh at five sites in each lake. Sites were chosen to represent a variety of habitat types and based on ability to effectively use the seine. Seining sites remained fixed for the duration of the study. Seines were used during daylight hours on each lake. Catch per unit effort was calculated as the number of individuals per seine haul. When water temperatures fell below 21°C (early September), we sampled age-0 walleyes using nighttime boat electrofishing (Wisconsin‐style; AC; 2.0–3.0 amps, 200–350 V, 25% duty cycle, two netters). The entire shoreline of each lake was sampled. Surveys were conducted prior to walleye fingerling stocking. All collected walleye were measured (TL, mm). Catch per unit effort was calculated as the number of age-0 walleyes per meter shoreline.
Removal efforts
In addition to standardized surveys in our experimental lake, in 2018 we began centrarchid removal efforts using a variety of techniques including fyke nets, boat electrofishing, mini-fyke nets, and cloverleaf traps (Fig. 1). Following annual spring fyke net surveys, fyke nets remained in the experimental lake to remove centrarchids. In 2018, we sampled 10 fyke nets from May 14 to June 7 when centrarchid catches started to decline. Due to personnel limitations, in 2019 and 2020 only five fyke nets were used from late spring (May 9, April 30) until late June (June 27, June 25). Additionally, we sampled five mini-fyke nets and 21 cloverleaf traps from late May through mid-August. All gears were emptied every 1-2 days and sites were rotated to maximize centrarchid catches. Collected fish were identified to species and measured (TL, mm). Centrarchid species were retained while other species were returned to McDermott Lake.

Methods

We sampled non-zebra mussel benthic macroinvertebrates twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along each of three transects (A-C) running perpendicular to the shore of Lake Mendota. We collected triplicate samples from each site using a 0.625 m-2 circular quadrat and an airlift method with a modified SCUBA tank suction device called an AquaVac. Air was released through a PVC pipe, creating backpressure to lift sediment, which was captured in a 500μm mesh bag and transported in a resealable plastic bag. We chose an airlift method because of difficulty closing Eckman samplers on the hard substrates of rock and zebra mussel druses. Occasionally additional samples were taken with an Eckman, often at deeper depths, for comparing to the main transects and depths sampled with AquaVac or to collect additional material for isotope analysis.<br/>We sampled non-zebra mussel benthic macroinvertebrates twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along each of three transects (A-C) running perpendicular to the shore of Lake Mendota. We collected triplicate samples from each site using a 0.625 m-2 circular quadrat and an airlift method with a modified SCUBA tank suction device called an AquaVac. Air was released through a PVC pipe, creating backpressure to lift sediment, which was captured in a 500μm mesh bag and transported in a resealable plastic bag. We chose an airlift method because of difficulty closing Eckman samplers on the hard substrates of rock and zebra mussel druses. Occasionally additional samples were taken with an Eckman, often at deeper depths, for comparing to the main transects and depths sampled with AquaVac or to collect additional material for isotope analysis.<br/>

Methods

We sampled adult zebra mussels twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along each of three transects running perpendicular to shore (A-C). Dominant substrates at transect A were rock at 1 m depth, sand at 3 and 5 m, and muck at 8 and 10 m. At transects B and C, sand was the dominant substrate at 1 and 3 m depth and muck was dominant at 5, 8, and 10m. Significant macrophyte growth was generally absent at all sites in June and occurred mostly at 1, 3, and 5 m sites only at transects A and C. Because most sites lacked hard substrate (rocks, logs, etc.) suitable for zebra mussel colonization, we also sampled five additional rocky 1m depth sites to represent prime zebra mussel habitat.<br/>We sampled adult zebra mussels twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along each of three transects running perpendicular to shore (A-C). Dominant substrates at transect A were rock at 1 m depth, sand at 3 and 5 m, and muck at 8 and 10 m. At transects B and C, sand was the dominant substrate at 1 and 3 m depth and muck was dominant at 5, 8, and 10m. Significant macrophyte growth was generally absent at all sites in June and occurred mostly at 1, 3, and 5 m sites only at transects A and C. Because most sites lacked hard substrate (rocks, logs, etc.) suitable for zebra mussel colonization, we also sampled five additional rocky 1m depth sites to represent prime zebra mussel habitat.<br/>We sampled adult zebra mussels twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along each of three transects running perpendicular to shore (A-C). Dominant substrates at transect A were rock at 1 m depth, sand at 3 and 5 m, and muck at 8 and 10 m. At transects B and C, sand was the dominant substrate at 1 and 3 m depth and muck was dominant at 5, 8, and 10m. Significant macrophyte growth was generally absent at all sites in June and occurred mostly at 1, 3, and 5 m sites only at transects A and C. Because most sites lacked hard substrate (rocks, logs, etc.) suitable for zebra mussel colonization, we also sampled five additional rocky 1m depth sites to represent prime zebra mussel habitat.

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

We sampled phytobenthos twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along three transects running perpendicular to shore (A-C, Fig. 1). We collected triplicate samples at each site. SCUBA divers retrieved one rock at rock-dominated sites, or a petri dish full of undisturbed sediment at sand- and muck-dominated sites, and transported samples to the surface in a resealable plastic bag. In the laboratory, we scrubbed phytobenthos from rocks with a brush or emptied petri dish contents into a beaker. We separated phytobenthos from inorganic material by adding ~1 L of deionized water, homogenizing the sample, allowing settlement of inorganic material, and decanting the suspended phytobenthos. We kept samples dark and refrigerated until completely processed to prevent cell division after collection. <br/>We sampled phytobenthos twice a summer (early June and late August) from 2016-2018 at five depths (1, 3, 5, 8, and 10m) along three transects running perpendicular to shore (A-C, Fig. 1). We collected triplicate samples at each site. SCUBA divers retrieved one rock at rock-dominated sites, or a petri dish full of undisturbed sediment at sand- and muck-dominated sites, and transported samples to the surface in a resealable plastic bag. In the laboratory, we scrubbed phytobenthos from rocks with a brush or emptied petri dish contents into a beaker. We separated phytobenthos from inorganic material by adding ~1 L of deionized water, homogenizing the sample, allowing settlement of inorganic material, and decanting the suspended phytobenthos. We kept samples dark and refrigerated until completely processed to prevent cell division after collection.

Methods

At each study site, teams visually surveyed the area for signs of
jumping worm presence, including live organisms or the characteristic
granular soil signature indicative of their activity. For example, in
a residential yard, participants would walk through the space for
approximately 10 minutes, brushing aside leaf litter and checking
underneath planters or landscaping cloth (where the species are
anecdotally known to congregate) for live earthworms, and examining
garden soil for structural characteristics. Next, earthworms were
censused at three haphazard locations using a 30cm x 30cm quadrat and
a standard mustard extraction (Lawrence and Bowers 2002). Any
suspected jumping worms found were collected and returned to the
laboratory for visual identification following the field campaign. We
identified jumping worms to species (A. tokioensis, A. agrestis, M.
hilgendorfi) when possible (Chang et al. 2016a). Participants also
recorded the presence/absence of any additional (European) earthworm
species observed during sampling.
<br/>