Methods

Sampling methods are described by Szydlowski et al. "Three decades of lake
monitoring reveals community recovery after population declines of invasive rusty crayfish
(Faxonius rusticus)," but are provided here for convenience. Instrumentation is further
documented in the supplementary information of the paper. Macrophytes were sampled during
July and August at a subset of crayfish sampling sites within our ten study lakes (n =
6–14 sites per lake) that were selected in 1987 to capture a variety of substrates and
both east and west sun exposure. Sampling depths were randomly assigned to sites during
initial sampling in 1987 as either 0.75 m, ½ of Secchi depth, or ¾ of Secchi depth, with
1987 Secchi depths used for all subsequent sampling years for macrophyte surveys. We
followed the line-intercept method to sample macrophytes, using snorkeling and SCUBA to
visually identify and determine the presence or absence of macrophyte species along a 25 m
transect set parallel to shore at the pre-determined depth for each sampling site.
Transects were marked at 1 m intervals, with the first 10 cm of each interval marked by a
band of tape. Divers moved along the transect recording the presence or absence of each
macrophyte species crossing the vertical plane of each 10 cm band. The line-intercept
method allowed us to obtain a measure of both macrophyte species richness and abundance.
Because just presence or absence of macrophyte species was recorded, and only at each 10
cm band, our measurements provide an index for abundance and a minimum estimate for
species richness. Freshwater snails were sampled at locations historically sampled for
crayfish (n = 24 or 36 sites per lake) between late June and early August. As with
macrophytes, snails were sampled at randomly assigned depths of either 0.75 m, ½ of Secchi
depth, or ¾ of Secchi depth. While the same absolute depths were used in 1987 and 2002
based on 1987 Secchi values, depths in 2011 and 2020 were determined using year-specific
Secchi values. Most sampling depths in 2011 and 2020 varied only slightly from the 1987
and 2002 values, but in two lakes the change in sampling depth was greater than one meter
due to larger shifts in water clarity. The greatest changes in sampling depth (2.7 m in
Papoose Lake and 1.5 m in Little John Lake) occurred at the ¾ Secchi depth sites, whereas
the ½ Secchi depth sites were less affected by the change in water clarity in these two
lakes. We sampled snails using methods and equipment designed for each habitat type
present in our study lakes (soft substrates, macrophytes, and cobble). For soft substrates
such as sand and muck (flocculent sediment or sediment rich in organic material), we used
a cylindrical polyvinyl chloride (PVC) sediment corer (0.018 m2), which we used to take a
5 cm sediment core. For sites with soft substrates where macrophytes were present, we used
a modified PVC sampler of the same size but with two hinged PVC halves, and a net made of
1-mm mesh attached to the top. We carefully closed the two halves of the PVC sampler
around macrophytes growing at the surface and zippered the mesh net around taller
macrophytes before pushing the corer into the sediment to collect a 5 cm core. Collecting
the macrophyte material along with the sediment allowed us to sample any snails on the
macrophytes along with those in the sediment. At the water’s surface we sieved (with 1 mm
mesh) all cores from soft substrates to remove fine sediments and large particles and
picked through macrophyte material for snails. Finally, for cobble habitats, we placed a
ring (0.1 or 0.5 m2) on the substrate at each site to define a sampling area. In 1987 and
2002, the 0.1 m2 ring was used for sites with a high density of snails, and the 0.5 m2
ring was used for sites with a low density of snails. In 2011 and 2020, we used the 0.5 m2
ring at all sites. We gently collected the surface layer of rocks within the sampling ring
and briefly brought the rocks to the surface, where we scraped attached material into a
collection pan and funneled it through a 1 mm mesh sieve to gather snails. We stored
snails collected using all sampling methods in 70% ethanol for later identification. In
the lab, we picked snails from all samples and identified them to species or genus (for
Physella sp.) according to Burch (1989) and Johnson et al. (2013), with revisions for
Lymnaeidae (Hubendick 1951) and Planorbidae (Hubendick and Rees 1955). We calculated snail
abundance as density to account for differences between the sediment corers and the rings
in area sampled. Snail samples from 1987 were lost in a laboratory flood, but specimens
from 2002 and 2011 are vouchered at the Notre Dame Museum of Biodiversity in Notre Dame,
Indiana, USA. Specimens from 2020 are vouchered at the Illinois Natural History Survey
Mollusk Collection at the University of Illinois in Champaign, Illinois, USA. In 2020, we
were not able to sample macrophytes and snails using SCUBA due to limitations from the
COVID-19 pandemic. Therefore, we excluded a small portion of deeper sites (approximately
2% of total macrophyte sites and 13% of total snail sites) that could not be sampled
accurately and safely while snorkeling. In addition, because of a few lost samples, data
from previous sampling years were not always available for each site. Consequently, in our
datasets of macrophytes and snails, we only include sites for which we had data in all
four sampling years (n = 100 sites/year for macrophytes, n = 208 sites/year for snails).
In our snail data, we only included snails which were alive at the time of sampling (i.e.,
we did not include empty shells).<br/>

Methods

Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>Boaters were recruited into a yearlong trip diary program at the landings of lakes in Vilas and Oneida Counties (Wisconsin) between Memorial Day weekend and Labor Day, 2011.<br/>

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

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/>

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/>

Methods

Two 30-meter vertical tows (0.5m diameter, 400um mesh net) are collected at the deepest part of Trout Lake each time the lake is visited for routine LTER sampling during open water. On occasion, tows are collected on additional dates. Samples are visually scanned in their entirety for number of Bythotrephes present. The samples are not preserved or archived.

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”.