US Long-Term Ecological Research Network

LTREB Biological Limnology at Lake Myvatn 2012-current

Abstract
These data are part of a long-term monitoring program in the central part of Myvatn that represents the dominant habitat, with benthos consisting of diatomaceous ooze. The program was designed to characterize import benthis and pelagic variables across years as midge populations varied in abundance. Starting in 2012 samples were taken at roughly weekly inervals during June, July, and August, which corresponds to the summer generation of the dominant midge,<em>Tanytarsus gracilentus</em>.
Creator
Dataset ID
296
Date Range
-
Maintenance
Ongoing
Metadata Provider
Methods
Benthic Chlorophyll Field sampling (5 samples) (2012, 2013)1. Take 5 cores from the lake2. Cut the first 0.75 cm (1 chip) of the core with the extruder and place in deli container. Label with date and core number.3. Place deli containers into opaque container (cooler) and return to lab. This is the same sample that is used for the organic matter analysis.In 2014, the method for sampling benthic chlorophyll changed. The calculation of chlorophyll was changed to reflect the different area sampled. Below is the pertinent section from the methods protocols. Processing after the collection of the sample was not changed.Take sediment samples from the 5 cores collected for sediment characteristics. Take 4 syringes of sediment with 10mL syringe (15.96mm diameter). Take 4-5cm of sediment. Then, remove bottom 2cm and place top 2cm in the film canister.Filtering1. Measure volume of material in deli container with 60mL syringe and record.2. Homogenize and take 1mL sample with micropipette. The tip on the micropipette should be cut to avoid clogging with diatoms. Place the 1mL sample in a labeled film canister. Freeze sample at negative 20 degrees Celsius unless starting methanol extraction immediately.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec.4. After 6-18 hours, shake container for 5 sec.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 per cent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000 microLiter pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120microLiters of 0.1 N HCl (30microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Chlorophyll Field sampling (5 samples)1. Take 2 samples at each of three depths, 1, 2, and 3m with Arni&rsquo;s zooplankton trap. For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. 2. Empty into bucket by opening the bottom flap with your hand.3. Take bucket to lab.Filtering1. Filter 1L water from integrated water sample (or until the filter is clogged) through the 47 mm GF/F filter. The pressure used during filtering should be low ( less than 5 mm Hg) to prevent cell breakage. Filtering and handling of filters should be performed under dimmed lighting.2. Remove the filter with forceps, fold it in half (pigment side in), and put it in the film canister. Take care to not touch the pigments with the forceps.3. Add 20mL methanol. This methanol can be kept cool in the fridge, although then you will need a second bottle of methanol for the fluorometer. Shake for 5 sec. and place in fridge.4. After 6-18 hours, shake container for 5 sec.5. Analyze sample in fluorometer after 24 hours.Fluorometer1. Allow the film canisters to sit at room temperature for approximately 15 min to avoid excessive condensation on the glass tubes. Shake tubes for 5 sec after removing from fridge but then be careful to let them settle before removing sample.2. Record the sample information for all of the film canisters on the data sheet.3. Add 4mL of sample to a 13x100mL glass tube.4. Insert the sample into the fluorometer and record the reading in the Fluor Before Acid column. The sample reading should be close to one of the secondary solid standards (42ug/L or 230ug/L), if not, dilute the sample to within 25 percent of the secondary solid standards (30-54ug/L or 180-280ug/L). It is a good idea to quickly check 2mL of a sample that is suspected to be too high to get an idea if other samples may need to be diluted. If possible, read the samples undiluted.5. If a sample needs to be diluted, use a 1000uL pipette and add 2mL of methanol to a tube followed by 2mL of undiluted sample. Gently invert the tube twice and clean the bottom with a paper towel before inserting it into the fluorometer. If the sample is still outside of the ranges above, combine 1 mL of undiluted sample with 3 mL of methanol. Be sure to record the dilution information on the data sheet.6. Acidify the sample by adding 120 microLiters of 0.1 N HCl (30 microLiters for every one mL of sample). Then gently invert the sample and wait 90 seconds (we used 60 seconds in 2012, the protocol said 90) before putting the sample into the fluorometer and recording the reading in the Fluor After Acid column. Be sure to have acid in each tube for exactly the same amount of time. This means doing one tube at a time or spacing them 30-60 seconds apart.7. Double check the results and redo samples, which have suspicious numbers. Make sure that the after-acidification values make sense when compared to the before acidification value (the before acid/after acid ratio should be approximately the same for all samples).Clean up1. Methanol can be disposed of down the drain as long as at least 50 times as much water is flushed.2. Rinse the film canisters and lids well with tap water and scrub them out with a bottle brush making sure to remove any remaining filter paper. Give a final rinse with distilled water. Pelagic Zooplankton Counts Field samplingUse Arni&rsquo;s zooplankton trap (modified Schindler) to take 2 samples at each of 1, 2, and 3m (6 total). For the 1m sample, drop the trap to the top of the chain. Each trap contains about 2.5L of water when full. Integrate samples in bucket and bring back to lab for further processing.Sample preparation in lab1. Sieve integrated plankton tows through 63&micro;m mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micro meter mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted as well.6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Benthic Microcrustacean Counts Field samplingLeave benthic zooplankton sampler for 24h. Benthic sampler consists of 10 inverted jars with funnel traps in metal grid with 4 feet. Set up on bench using feet (on side) to get a uniform height of the collection jars (lip of jar = 5cm above frame). Upon collection, pull sampler STRAIGHT up, remove jars, homogenize in bucket and bring back to lab. Move the boat slightly to avoid placing sampler directly over cored sediment.Sample preparation in lab1. Sieve integrated samples through 63 micrometer mesh and record volume of full sample2. Collect in Nalgene bottles and make total volume to 50mL3. Add 8 drops of lugol to fix zooplankton.4. Label bottle with sample date, benthic or pelagic zooplankton, and total volume sieved. Samples can be stored in the fridge until time of countingCounting1. Remove sample from fridge2. Sieve sample with 63 micrometer mesh over lab sink to remove Lugol&rsquo;s solution (which vaporizes under light)3. Suspend sample in water in sieve and flush from the back with squirt bottle into counting tray4. Homogenize sample with forceps or plastic pipette with tip cut off5. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too!6. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridgeSubsamplingIf homogenized original sample contains more than 500 individuals in the first line of counting tray, you may subsample under the following procedure.1. Return original sample to Nalgene bottle and add water to 50mL2. Homogenize sample by swirling Nalgene bottle3. Collect 10mL of zooplankton sample with Hensen-Stempel pipette4. Empty contents of Hensen-Stempel pipette into large Bogorov tray5. Homogenize sample in tray with forceps or plastic pipette with tip cut off6. Identify (see zooplankton identification guide) using backlit microscope and count with multiple-tally counter. i. Set magnification so that you can see both top and bottom walls of the tray. ii. Change focus depth to check for floating zooplankton that must be counted, too! 7. Pipette sample back into Nalgene bottle, add water to 50mL, add 8 drops Lugol&rsquo;s solution, and return to fridge Chironomid Counts (2012, 2013) For first instar chironomids in top 1.5cm of sediment only (5 samples)1. Use sink hose to sieve sediment through 63 micrometer mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents into small deli container.3. Return label to deli cup (sticking to underside of lid works well).For later instar chironomids in the section 1.5-11.5cm (5 samples)4. Sieve with 125 micrometer mesh in the field.5. Sieve through 125micrometer mesh again in lab to reduce volume of sample.6. Transfer sample to deli container or pitfall counting tray.For all chironomid samples7. Under dissecting scope, pick through sieved contents for midge larvae. You may have to open tubes with forceps in order to check for larvae inside.8. Remove larvae with forceps while counting, and place into a vial containing 70 percent ethanol. Larvae will eventually be sorted into taxonomic groups (see key). You may sort them into taxonomic groups as you pick the larvae, or you can identify the larvae while measuring head capsules if chironomid densities are low (under 50 individuals per taxanomic group).9. For a random sample of up to 50 individuals of each taxonomic group, measure head capsule, see Chironomid size (head capsule width).10. Archive samples from each sampling date together in a single 20mL glass vial with screw cap in 70 percent ethanol and label with sample contents , Chir, sample date, lake ID, station ID, and number of cores. Chironomid Cound (2014) In 2014, the method for sampling chironomid larvae changed starting with the sample on 2014-06-27; the variable &quot;top_bottom&quot; is coded as a 2. In contrast to previous measurements, the top and bottom core samples were combined and then subsampled. Below is the pertinent section of the protocols.Chironomid samples should be counted within 24 hours of collection. This ensures that larvae are as active and easily identified as possible, and also prevents predatory chironomids from consuming other larvae. Samples should be refrigerated upon returning from the field.<strong>For first instar chironomids in top 1.5cm of sediment only (5 samples)</strong>1. Use sink hose to sieve sediment through 63&micro;m mesh. You may use moderate pressure to break up tubes.2. Back flush sieve contents using a water bottle into small deli container.3. Return label to deli cup (sticking to underside of lid works well).<strong>For larger instar chironomids in the section 1.5-11.5cm (5 samples)</strong>4. Sieve with 125&micro;m mesh in the field.5. Sieve through 125&micro;m mesh again in lab to reduce volume of sample and break up tubes.6. Transfer sample to deli container with the appropriate label.<strong>Subsample if necessary</strong>If necessary, subsample with the following protocol.a. Combine top and bottom samples from each core (1-5) in midge sample splitter.b. Homogenize sample thoroughly, collect one half in deli container, and label container with core number and &ldquo;1/2&rdquo;c. If necessary, split the half that remains in the sampler into quarters, and collect each in deli containers labeled with core number, &ldquo;1/4&rdquo;, and replicate 1 or 2d. Store all deli containers in fridge until counted, and save until all counting is complete&quot; Chironomid Size (head capsule width) 1. Obtain picked samples preserved in ethanol and empty onto petri dish.2. Sort larvae by family groups, arranging in same orientation for easy measurment.3. Set magnification to 20, diopter, x 50 times4. Take measurments for up to 50 or more individuals of each taxa. Round to nearest optical micrometer unit.5. Fill out data sheet for number of larvae in each taxa, Chironomid measurements for each taxa, date of sample, station sample was taken from, which core the sample came from, who picked the core, and your name as the measurer.6. Enter data into shared sheetSee &quot;Chironomid Counts&quot; for changes in sampling chironomid larvae in 2014.
Version Number
17

Fluxes project at North Temperate Lakes LTER: Spatial Metabolism Study 2007

Abstract
Data from a lake spatial metabolism study by Matthew C. Van de Bogert for his Phd project, "Aquatic ecosystem carbon cycling: From individual lakes to the landscape."; The goal of this study was to capture the spatial heterogeneity of within-lake processes in effort to make robust estimates of daily metabolism metrics such as gross primary production (GPP), respiration (R), and net ecosystem production (NEP). In pursuing this goal, multiple sondes were placed at different locations and depths within two stratified Northern Temperate Lakes, Sparkling Lake (n=35 sondes) and Peter Lake (n=27 sondes), located in the Northern Highlands Lake District of Wisconsin and the Upper Peninsula of Michigan, respectively.Dissolved oxygen and temperature measurements were made every 10 minutes over a 10 day period for each lake in July and August of 2007. Dissolved oxygen measurements were corrected for drift. In addition, conductivity, temperature compensated specific conductivity, pH, and oxidation reduction potential were measured by a subset of sondes in each lake. Two data tables list the spatial information regarding sonde placement in each lake, and a single data table lists information about the sondes (manufacturer, model, serial number etc.). Documentation :Van de Bogert, M.C., 2011. Aquatic ecosystem carbon cycling: From individual lakes to the landscape. ProQuest Dissertations and Theses. The University of Wisconsin - Madison, United States -- Wisconsin, p. 156. Also see Van de Bogert, M.C., Bade, D.L., Carpenter, S.R., Cole, J.J., Pace, M.L., Hanson, P.C., Langman, O.C., 2012. Spatial heterogeneity strongly affects estimates of ecosystem metabolism in two north temperate lakes. Limnology and Oceanography 57, 1689-1700.
Core Areas
Dataset ID
285
Date Range
-
Metadata Provider
Methods
Data were collected from two lakes, Sparkling Lake (46.008, -89.701) and Peter Lake (46.253, -89.504), both located in the northern highlands Lake District of Wisconsin and the Upper Peninsula of Michigan over a 10 day period on each lake in July and August of 2007. Refer to Van de Bogert et al. 2011 for limnological characteristics of the study lakes.Measurements of dissolved oxygen and temperature were made every 10 minutes using multiple sondes dispersed horizontally throughout the mixed-layer in the two lakes (n=35 sondes for Sparkling Lake and n=27 sondes for Peter Lake). Dissolved oxygen measurements were corrected for drift.Conductivity, temperature compensated specific conductivity, pH, and oxidation reduction potential were also measured by a subset of sensors in each lake. Of the 35 sondes in Sparkling Lake, 31 were from YSI Incorporated: 15 of model 600XLM, 14 of model 6920, and 2 of model 6600). The remaining sondes placed in Sparkling Lake were 4 D-Opto sensors, Zebra-Tech, LTD. In Peter Lake, 14 YSI model 6920 and 13 YSI model 600XLM sondes were used.Sampling locations were stratified randomly so that a variety of water depths were represented, however, a higher density of sensors were placed in the littoral rather than pelagic zone. See Van de Bogert et al. 2012 for the thermal (stratification) profile of Sparkling Lake and Peter Lake during the period of observation, and for details on how locations were classified as littoral or pelagic. In Sparkling Lake, 11 sensors were placed within the shallowest zone, 12 in the off-shore littoral, and 6 in each of the remaining two zones, for a total of 23 littoral and 12 pelagic sensors. Similarly, 15 sensors were placed in the two littoral zones, and 12 sensors in the pelagic zone.Sensors were randomly assigned locations within each of the zones using rasterized bathymetric maps of the lakes and a random number generator in Matlab. Within each lake, one pelagic sensor was placed at the deep hole which is used for routine-long term sampling.Note that in Sparkling Lake this corresponds to the location of the long-term monitoring buoy. After locations were determined, sensors were randomly assigned to each location with the exception of the four D-Opto sensor is Sparkling Lake, which are a part of larger monitoring buoys used in the NTL-LTER program. One of these was located near the deep hole of the lake while the other three were assigned to random locations along the north shore, south shore and pelagic regions of the lake. Documentation: Van de Bogert, M.C., Bade, D.L., Carpenter, S.R., Cole, J.J., Pace, M.L., Hanson, P.C., Langman, O.C., 2012. Spatial heterogeneity strongly affects estimates of ecosystem metabolism in two north temperate lakes. Limnology and Oceanography 57, 1689-1700.
Version Number
17

Lake Metabolism

Study sites
We sampled surface waters of 31 lakes in the Northern Highland Lake district of Wisconsin and the Upper Peninsula of Michigan during July and August of 2000 (Table 1). The lakes were chosen to span wide and orthogonal ranges in DOC and TP concentrations and for their close proximity to the Trout Lake Station in Vilas county, Wisconsin. The order in which the lakes were sampled was randomized.

North Temperate Lakes LTER: Physical Limnology of Primary Study Lakes 1981 - current

Abstract
Parameters characterizing the physical limnology of the eleven primary lakes (Allequash, Big Muskellunge, Crystal, Sparkling, Trout, bog lakes 27-02 [Crystal Bog] and 12-15 [Trout Bog], Mendota, Monona, Wingra and Fish) are measured at one station in the deepest part of each lake at 0.25-m to 1-m depth intervals depending on the lake. Measured parameters in the data set include water temperature, vertical penetration of photosynthetically active radiation (PAR; not measured on lakes Mendota, Monona, Wingra, and Fish), dissolved oxygen, as well as the derived parameter percent oxygen saturation. Sampling Frequency: fortnightly during ice-free season - every 6 weeks during ice-covered season for the northern lakes. The southern lakes are similar except that sampling occurs monthly during the fall and typically only once during the winter (depending on ice conditions). Number of sites: 11
Core Areas
Dataset ID
29
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
Light (PAR) extinction coefficient is calculated by linearly regressing ln (FRLIGHT (z)) on depth z where the intercept is not constrained. FRLIGHT(z) = LIGHT(z) or DECK(z) where LIGHT(z) is light measured at depth z and DECK(z) is light measured on deck (above water) at the same time. For open water light profiles, the surface light measurement (depth z = 0) is excluded from the regression. For winter light profiles taken beneath the ice, the first light data are taken at the bottom of the ice cover and are included in the regression. The depth of uppermost light value is equal to the depth of the ice adjusted by the water level in the sample hole, i.e., the depth below the surface of the water. The water level can be at, above or below the surface of the ice. If the water level was not recorded, it is assumed to be 0.0 and the calculated light extinction coefficient is flagged. If ice thickness was not recorded, a light extinction coefficient is not calculated. For light data collected prior to March, 2007, light values less than 3.0 (micromolesPerMeterSquaredPerSec) are excluded. For light data collected starting in March 2007, light values less than 1.0 (micromolesPerMeterSquaredPerSec) are excluded. Except for bog lakes before August 1989, a light extinction coefficient is not calculated if there are less than three FRLIGHT values to be regressed. For bog lakes before August 1989, a light extinction coefficient is calculated if there are least two FRLIGHT values to be regressed. In these cases, the light extinction coefficient is flagged as non-standard. FRLIGHT values should be monotonically decreasing with depth. For light profiles where this is not true, a light extinction coefficient is not calculated. For samples for which light data at depth are present, but the corresponding deck light are missing, a light extinction coefficient is calculated by regressing ln (LIGHT (z)) on depth z. Note that if actual deck light had remained constant during the recording of the light profile, the resulting light extinction coefficient is the same as from regressing ln(FRLIGHT(z)). In these cases, the light extinction coefficient is flagged as non-standard. Oxygen and Temperature: We sample at the deepest part of the lake, taking a temperature and oxygen profile at meter intervals from the surface to within 1 meter of the bottom using a YSI Pro-ODO temporDO meter. We sample biweekly during open water and approximately every five weeks during ice cover. Protocol Log: Prior to 2011, we used a YSI Model 58 temporDO meter.
Short Name
NTLPH01
Version Number
27

North Temperate Lakes LTER: High Frequency Water Temperature Data - Sparkling Lake Raft 1989 - current

Abstract
The instrumented raft on Sparkling Lake is equipped with a thermistor chain that measures water temperature from depths ranging from the surface to 18m at intervals from 0.5 to 3m throughout the water column. The surface temperature sensors are attached to floats so that they are as close to the surface as feasible. The raft on Sparkling Lake is also equipped with a dissolved oxygen sensor and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, evaporation rates, and lake metabolism. Estimating the flux of solutes to and from lakes requires accurate water budgets. Evaporation rates are a critical component of the water budget of lakes. Data from the instrumented raft on Sparkling Lake includes micrometeorological parameters from which evaporation can be calculated. Raft measurements of relative humidity and air temperature (2 m height), wind velocity (1, 2, and 3 m heights), and water temperatures are combined with measurements of total long-wave and short-wave radiation data from a nearby shore station to determine evaporation by the energy budget technique. Comparable evaporation estimates from mass transfer techniques are calibrated against energy budget estimates to produce a lake-specific mass transfer coefficient for use in estimating evaporation rates Sampling Frequency: one minute; averaged to hourly and daily values as well as higher resolution values such as 2 min and 10 min. Number of sites: 1
Dataset ID
5
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
see abstract for methods description
Short Name
NTLEV02
Version Number
21

North Temperate Lakes LTER: High Frequency Meteorological and Dissolved Oxygen Data - Sparkling Lake Raft 1989 - current

Abstract
The instrumented raft on Sparkling Lake is equipped with a dissolved oxygen and CO2 sensors, a thermistor chain, and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, evaporation rates, and lake metabolism. Estimating the flux of solutes to and from lakes requires accurate water budgets. Evaporation rates are a critical component of the water budget of lakes. Data from the instrumented raft on Sparkling Lake includes micrometeorological parameters from which evaporation can be calculated. Raft measurements of relative humidity and air temperature (2 m height), wind velocity ( at 1, 2, and 3 m heights; but beginning in 2008, only at 2 m) ,and water temperatures (from thermistors placed throughout the water column at intervals varying from 0.5 to 3m) are combined with measurements of total long-wave and short-wave radiation data from a nearby shore station to determine evaporation by the energy budget technique. Comparable evaporation estimates from mass transfer techniques are calibrated against energy budget estimates to produce a lake-specific mass transfer coefficient for use in estimating evaporation rates. After correcting for flux to or from the atmosphere and vertical mixing within the water column, high frequency measurements of dissolved gases such as carbon dioxide and oxygen can be used to estimate gross primary productivity, respiration, and net ecosystem productivity, the basic components of whole lake metabolism. Other parameters measured include precipitation, wind direction (beginning in 2008), and barometric pressure (beginning in 2008). Sampling Frequency: one minute; averaged to hourly and daily values as well as higher resolution values such as 2 min and 10 min. Number of sites: 1
Core Areas
Dataset ID
4
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
The instrumented raft on Sparkling Lake is equipped with a D-Opto dissolved oxygen sensor, a thermistor chain, and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, evaporation rates, and lake metabolism. Estimating the flux of solutes to and from lakes requires accurate water budgets. Evaporation rates are a critical component of the water budget of lakes. Data from the instrumented raft on Sparkling Lake includes micrometeorological parameters from which evaporation can be calculated. Raft measurements of relative humidity and air temperature (2 m height), wind velocity ( at 1, 2, and 3 m heights; but beginning in 2008, only at 2 m) ,and water temperatures (from thermistors placed throughout the water column at intervals varying from 0.5 to 3m) are combined with measurements of total long-wave and short-wave radiation data from a nearby shore station to determine evaporation by the energy budget technique. Comparable evaporation estimates from mass transfer techniques are calibrated against energy budget estimates to produce a lake-specific mass transfer coefficient for use in estimating evaporation rates. After correcting for flux to or from the atmosphere and vertical mixing within the water column, high frequency measurements of dissolved gases such as carbon dioxide and oxygen can be used to estimate gross primary productivity, respiration, and net ecosystem productivity, the basic components of whole lake metabolism. Other parameters measured include precipitation, wind direction (beginning in 2008), and barometric pressure (beginning in 2008). Sampling Frequency: one minute; averaged to hourly and daily values as well as higher resolution values such as 2 min and 10 min.Dissolved oxygen sensors: 2004-2006: Greenspan Technology series 1200; 2007-2016: Zebra-Tech Ltd. D-Opto; 2018+: OTT HydrolabCO2 sensors: 2018+: ProOceanos MiniCO2 for dissolved CO2; Eosense Inc. eosGP for atmospheric CO2
Short Name
NTLEV01
Version Number
32

North Temperate Lakes LTER: High Frequency Water Temperature Data - Sparkling Bog North Buoy 2008 - current

Abstract
The instrumented buoy on Sparkling Bog North is equipped with a thermistor chain that measures water temperature at the surface, at 0.25 m and at every .5 m from 0.5 m to 4.5 m. The surface temperature sensors are attached to floats so that they are as close to the surface as feasible. The buoy is also equipped with a dissolved oxygen sensor, meteorological sensors, a CO2 sensor and a YSI AutoProfiler that provide fundamental information on lake thermal structure, weather conditions, and lake metabolism. Prior to May 2009, data were collected at 1 minute or 10 minute intervals. Since May 2009, data are being collected each minute. Hourly and daily water temperature averages are computed from high resolution data. Hourly and daily values may not be current with high resolution data. In 2008, the instrumented buoy was deployed in Sparkling Bog North from March 24 to November 10. In 2009, the buoy was deployed on the ice on March 7 and was not removed for the winter of 2009 to 2010. Sampling Frequency: varies for instantaneous sample. Generally 1 minute or 10 minutes. Number of sites: 1
Dataset ID
228
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
The instrumented buoy on Sparkling Bog North is equipped with a thermistor chain that measures water temperature at the surface, at 0.25 m and at every .5 m from 0.5 m to 4.5 m. The buoy is also equipped with a dissolved oxygen sensor, meteorological sensors, a CO2 sensor and a YSI AutoProfiler that provide fundamental information on lake thermal structure, weather conditions, and lake metabolism. Prior to May 2009, data were collected at 1 minute or 10 minute intervals. Since May 2009, data are being collected each minute. Hourly and daily water temperature averages are computed from high resolution data. Hourly and daily values may not be current with high resolution data. In 2008, the instrumented buoy was deployed in Sparkling Bog North from March 24 to November 10. In 2009, the buoy was deployed on the ice on March 7 and was not removed for the winter of 2009 to 2010. Sampling Frequency: varies for instantaneous sample.
Short Name
NSPBBUOY2
Version Number
18

North Temperate Lakes LTER: High Frequency Meteorological and Dissolved Oxygen Data - Sparkling Bog North Buoy 2008 - 2012

Abstract
The instrumented buoy on Sparkling Bog North is equipped with a dissolved oxygen sensor, a thermistor chain, and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, and lake metabolism. Data are usually collected either at 1 minute or 10 minute intervals. The D-Opto dissolved oxygen sensor is 0.5m from the lake surface, thermistors are at the surface, at 0.25 m and at every .5 m from 0.5 m to 4.5 m, and meteorological sensors measure wind speed, wind direction, relative humidity, and air temperature. The buoy is also equipped with a CO2 monitor and a YSI AutoProfiler that measures several parameters including dissolved oxygen, water temperature, conductivity, pH, ORP, turbulence and chlorophyll-a. After correcting for flux to or from the atmosphere and vertical mixing within the water column, high frequency measurements of dissolved gases such as carbon dioxide and oxygen can be used to estimate gross primary productivity, respiration, and net ecosystem productivity, the basic components of whole lake metabolism. Sampling Frequency: varies for instantaneous sample. Generally 1 minute or 10 minutes. Number of sites: 1
Core Areas
Dataset ID
227
Date Range
-
Maintenance
completed
Metadata Provider
Methods
see abstract for methods description
Short Name
NSPBBUOY1
Version Number
20

North Temperate Lakes LTER: High Frequency Water Temperature Data - Lake Mendota Buoy 2006 - current

Abstract
The instrumented buoy on Lake Mendota is equipped with a thermistor chain that measures water temperature. In 2006, the thermistors were placed every 0.5m from the surface through 7m and every 1m from 7m to 15m. In 2007and 2008, the thermistors were placed every 0.5 m from the surface through 2m and every 1m from 2m to 20m. The sensor at the water surface is as close to the surface as feasible. The sensor housing itself is ~5cm long, so we have it at about 1 to 6cm deep.The Lake Mendota buoy is also equipped with a dissolved oxygen sensor and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, and lake metabolism. Data are collected every minute. After correcting for flux to or from the atmosphere and vertical mixing within the water column, high frequency measurements of dissolved gases such as carbon dioxide and oxygen can be used to estimate gross primary productivity, respiration, and net ecosystem productivity, the basic components of whole lake metabolism. Hourly and daily water temperature averages are computed from high resolution (1 minute) data. Hourly and daily values may not be current with high resolution data.Deployment dates: 2006: June 27-Oct 17; 2007: May 18-Oct 22; 2008: June 26-Nov 4; 2009: April 17-Nov 23; 2010: April 9-Nov 10; 2011: May 19-Nov 11; 2012: April 30-Nov 30; 2013: April 27-Oct 27; 2014: May 4-Oct 12; 2015: June 25-Sept 1; 2016: March 29-Dec 12; 2017: April 23 - Nov 13; 2018: April 11-Nov 15; 2019: April 16-Nov 11Sampling Frequency: one minute. Number of sites: 1
Core Areas
Dataset ID
130
Date Range
-
Maintenance
ongoing
Metadata Provider
Methods
The instrumented buoy on Lake Mendota is equipped with a thermistor chain that measures water temperature. In 2006, the thermistors were placed every 0.5m from the surface through 7m and every 1m from 7m to 15m. In 2007and 2008, the thermistors were placed every 0.5 m from the surface through 2m and every 1m from 2m to 20m. The Lake Mendota buoy is also equipped with a dissolved oxygen sensor and meteorological sensors that provide fundamental information on lake thermal structure, weather conditions, and lake metabolism. Data are collected every minute. After correcting for flux to or from the atmosphere and vertical mixing within the water column, high frequency measurements of dissolved gases such as carbon dioxide and oxygen can be used to estimate gross primary productivity, respiration, and net ecosystem productivity, the basic components of whole lake metabolism. Hourly and daily water temperature averages are computed from high resolution (1 minute) data. Hourly and daily values may not be current with high resolution data.
Short Name
MEBUOY2
Version Number
29

Microbial Observatory at North Temperate Lakes LTER Alkaline Phosphatase Activity in Lake Mendota 2000 - 2001

Abstract
Parameters characterizing the alkaline phosphatase activity of Lake Mendota. Samples were collected at one station in the deepest part of each lake from an integrated sample of the epilimnion.Sampling Frequency: fortnightly during ice-free season - every 6 weeks during ice-covered seasonNumber of sites: 1
Core Areas
Dataset ID
45
Date Range
-
Metadata Provider
Methods
BEFOREHAND prepare MUF-P substrate. Dissolve 180.04 mg of MUF-P in 120.0 mL total fresh Nanopure water to get 5 mM solution. Aliquot 1 mL each into thirty microcentrifuge tubes. Store in freezer. These aliquots will be thawed as needed.BEFOREHAND prepare borate buffer by dissolving 4.7675 g in 250 mL Nanopure water.PREPARE MUF standards the day before and store in cold room. Standards are prepared as follows: Prepare 5 mM MUF by adding 158.56 mg MUF in 100 mL. Vortex. Dilute 0.120 mL of 5 mM MUF in 15.0 mL total to get 40 uM solution. Prepare solutions for working standards as follows. Vortex solutions before dilutions.MUF Concentration (nM) - Preparation:0 -- 0 mL of 200 nM in 10 mL.10 -- 0.5 mL of 200 nM in 10 mL.50 -- 2.5 mL of 200 nM in 10 mL.100 -- 5 mL of 200 nM in 10 mL.200 -- 75 mL of 200 nM in 10 mL.THAW MUF-P substrate the day before in cold room.LABEL scintillation vials day before sampling. For substrate need one vial labelled 100 uM. For lake water sample need 9 vials. Three replicates samples are run for each of three size fractions. The three size fractions are less than 0.2 um water, less than 1 um water and less than 70 um (field filtered) water.DAY of sampling, filter borate buffer solution through a 0.2 um syringe filter. Dilute 0.300 mL of 5 mM MUF-P solution in 15.0 mL total Nanopure water to get 100 uM solution. Vortex MUF-P before aliquoting to samples.TURN on fluorometer to warm up (10 minutes) and set filters to 365 nm excitation, 460 nm emmission. Aliquot 10 mL of lake water into appropriate 20 mL scintillation vials labelled the day before.PERFORM measurement in room with fluorometer in the dark. First run the standard curve. If the standard curve does not appear to be linear, make new standards before running samples. If standard curve is good, add 0.010 mL of 100 uM MUF-P substrate to each ten mL sample for a final concentration of 100 nM. Note time when you spiked the samples.BEGIN immediately aliquoting 3.0 mL of sample each to glass test tubes. Aliquot all nine test ubes before adding borate buffer. Add 1.0 mL borate buffer to test tubes and mix. Note fluorescence. Continue with next sample. Do less than70, less than1 and less than 0.2 um replicates together to make more comparable (i.e. less than70 um replicate 1, less than 1 um replicate 1, less than 0.2 um replicate 1, less than 70 um replicate 2, etc.). Note time when finisihed running all initial samples. Put rest of samples in incubator.REPEAT measurements of sample for final time point thirty to sixty minutes later. Note time when you begin aliquoting 3 mL of samples to test tubes. Note time when final samples have all be run.
NTL Themes
Short Name
MOACT1
Version Number
22
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